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biomni/tool/protocols/addgene/Addgene_ AAV Production.txt

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+PROTOCOL: AAV Production
+SOURCE: https://www.addgene.org/protocols/aav-production-hek293-cells/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+This protocol can be used to produce AAV from one Five Chambers Cell-Stack (CS5)
+(3,180 cm² - the same surface area as 21 x T-175 flasks). Cell stacks provide an
+efficient means to scale-up without having to handle a large number of T-175
+flasks.
+
+
+WORKFLOW TIMELINE
+================================================================================
+
+Day 0: Seed cells in CS2
+
+Day 2: Seed cells in CS5
+
+Day 3 (am): Transfect cells
+
+Day 7 (am): Harvest cells
+
+
+EQUIPMENT
+================================================================================
+
+• Class II, Type A2 Biological Safety Cabinet
+• 0.5–10 µL single channel pipette
+• 2–20 µL single channel pipette
+• 20–200 µL single channel pipette
+• 200–1000 µL single channel pipette
+• Hemocytometer or cell counter
+• Ice bucket
+• CO₂ incubator
+• Pipet controller
+• Hazardous waste container
+• pH meter
+• Stir plate
+• Magnetic stir bar
+• Sonicator
+• Ear protection
+• Vortex
+
+
+REAGENTS
+================================================================================
+
+• Adherent HEK293T cells (ideally AAV-293T clones)
+
+  💡 PRO TIP: While adherent, these cells are very loosely attached to the dish
+  surface and should be handled carefully. Avoid touching the cells when replacing
+  media.
+
+• T-175 flask, Corning 430825, 175 cm²
+• Cellstack 5, Corning 3319, 3180 cm²
+• Cellstack 2, Corning 3269, 1272 cm²
+• Heat-inactivated FBS (HI-FBS)
+
+  💡 PRO TIP: Different brands and lots of FBS can promote or inhibit
+  transfection. Test a variety of brands and lots of FBS to find one suitable for
+  your protocols. FBS can be purchased already heat-inactivated, or it can be
+  inactivated in the lab by heating to 56 °C for 30 minutes.
+
+• 0.45 μm polyethersulfone (PES) filter system, Nalgene, 565-0010 (or for larger
+  volumes 1000 mL 0.45 µm Rapid-Flow PES Filtration Unit, Nalgene 167-0045)
+
+  💡 PRO TIP: Do not use filters made of materials other than PES. AAV particles
+  stick to many other surfaces, but do not stick to PES. Using a PES filter will
+  maximize titer.
+
+• DMEM high glucose, Corning 10-013-CV
+• DMEM, low glucose (1 g/L) glucose, sodium pyruvate, Corning 10-014-CV (optional)
+• 7.5% sodium bicarbonate, Corning 25-035-CI (optional)
+• 1 M HEPES, HyClone SH30237.01 (optional)
+• L-alanyl-L-glutamine (or alternative stable glutamine such as glutaGRO,
+  Corning 25-015-CI)
+• Opti-MEM, Thermo Fisher 31985-070
+• 0.05% Trypsin/EDTA (e.g. TrypLE, Thermo Fisher, 12605010)
+• 1X PBS pH 7.4 without calcium or magnesium, Corning 21-040-CV (cations can
+  affect the attachment of adherent cells)
+• 1 mg/mL Polyethylenimine (PEI) 25 kDa MW
+
+  💡 PRO TIP: Other transfection reagents may be used in this protocol, but their
+  conditions must be optimized.
+
+• Plasmids for transfection: pHelper; pRC (Rep-Cap), plasmid expressing your gene
+  of interest
+• Triton X-100
+• Benzonase/DNAse I (Millipore 71205-3)
+• 40% Polyethylene Glycol 8000 (PEG) + 0.5 M NaCl
+• Cell lysis buffer (50 mM Tris HCl, 150 mM NaCl, 2 mM MgCl₂)
+
+
+REAGENT PREPARATION
+================================================================================
+
+DMEM Complete: 10% v/v FBS and 4 mM L-alanyl-L-glutamine (or stable alternative,
+such as glutaGRO)
+
+To a 500 mL bottle of DMEM high glucose, add 55 mL of heat-inactivated FBS and 5
+mL of glutaGRO 11 mL of 200 mM L-alanyl-L-glutamine. Store at 4 °C.
+
+D1 + 0.1 M sorbitol (optional): DMEM low glucose + 1% v/v HI-FBS + 0.1 M sorbitol
+
+In our hands, we have found that a lower glucose, lower FBS media causes less
+filter clogging during harvesting. The addition of sorbitol has been shown to
+increase viral titers by up to 1.8-fold (Rego et al., 2018).
+
+To prepare D1 + 0.1 M sorbitol, add 57 mL of 1.46 M sorbitol, 7.5 mL HI-FBS, and
+7.5 mL 100X glutaGRO, 7.5 mL of 7.5% Sodium Bicarbonate, 7.5 mL 1 M HEPES to 750
+mL DMEM + 1 g/L glucose.
+
+1 mg/mL polyethylenimine (PEI) solution:
+
+1. Dissolve 100 mg of PEI powder into 100 mL of deionized water.
+
+2. While stirring, slowly add hydrochloric acid until the solution clears.
+
+3. Check the pH of the solution and use hydrochloric acid or sodium hydroxide to
+   adjust the pH to 7.0.
+
+   💡 PRO TIP: The pH of this solution will drift pretty rapidly upon addition
+   of acid or base. Add only a few drops at a time. Allow them to mix and recheck
+   the pH to prevent over or undershooting the desired pH.
+
+4. Allow the solution to mix for 10 min and then recheck the pH to ensure that it
+   has not drifted.
+
+5. Filter the solution through a 0.22 μm membrane.
+
+6. Aliquot 500–1000 μL into sterile tubes.
+
+7. Store the tubes at -80 °C.
+
+8. After thawing, the solution can be stored at 4 °C for up to 2 months. After 2
+   months, discard the tube and thaw a new working stock.
+
+40% POLYETHYLENE GLYCOL (PEG) 8000 solution:
+
+1. Dissolve 400 g of PEG 8000 and 24 g of NaCl into deionized water and adjust to
+   a final volume of 1000 mL.
+
+2. Stir at RT until fully dissolved.
+
+   💡 PRO TIP: This step is challenging due to the high viscosity of PEG.
+   Stirring under medium heat will promote faster dissolution.
+
+3. Adjust the pH to ~7.4.
+
+4. Autoclave or sterile filter.
+
+   💡 PRO TIP: Stirring during the cooling period is recommended or the solution
+   may separate into phases.
+
+5. Aliquot and store at 4 °C.
+
+Cell Lysis Buffer: 50 mM Tris HCl, 150 mM NaCl, 2 mM MgCl₂
+
+1. Add the following to the 2 L sterile bottle: 1836 mL deionized water + 100 mL
+   of 1 M Tris HCl pH 8.5 + 60 mL of 5 M Sodium Chloride + 4 mL of 1 M Magnesium
+   Chloride
+
+2. Close the bottle and mix by inverting multiple times
+
+3. Adjust pH to ~8.5
+
+4. Filter sterilize through a 0.22 μm membrane.
+
+5. Store at 4 °C.
+
+
+CONSIDERATIONS BEFORE YOU START
+================================================================================
+
+• The health of the HEK293T cells is critical for optimal AAV yield.
+
+• Do not overgrow your cells. Pass the cells twice a week during the maintenance
+  phase and do not allow cells to reach 100% confluence (80–90% is ideal).
+
+• Pass and plate the cells the day before the transfection.
+
+• Thaw a new vial of cells after 30 passages.
+
+
+PROCEDURE
+================================================================================
+
+1. Trypsinize and resuspend the HEK293T cells from 2 x T-175 flasks. Cells should
+   be at ~80% confluence. For each T-175 flask:
+   • Aspirate culture media and rinse once with 10 mL of PBS.
+   • Aspirate PBS and add 2 mL of 0.05% Trypsin/EDTA. Wait ~2 min.
+   • Neutralize trypsin by adding 10 mL of DMEM Complete.
+   • Pipet back and forth vigorously multiple times to obtain a single cell
+     suspension (no clumps of cells).
+
+2. Pool cells from 2 x T-175 flasks. Adjust volume to 300 mL with DMEM complete
+   media and mix.
+
+3. Seed all cells in 1 CS2. Return to incubator for 48 h.
+
+4. Trypsinize and resuspend cells from the CS2. Cells should be at ~80%
+   confluence.
+   • Aspirate culture media and rinse once with 60 mL PBS.
+   • Aspirate PBS and add 35 mL of 0.05% Trypsin/EDTA.
+   • Wait ~2-3 minutes for cells to detach.
+   • Gently tap the sides of the CS2 to help detach the cells, add 200 mL of DMEM
+     complete media, then transfer the cells into a sterile bottle.
+   • Rinse the CS2 with 100 mL of DMEM complete medium and pool with the cells
+     harvested in the previous step.
+   • Pipette back and forth or shake vigorously to obtain a single cell suspension
+     (no clumps).
+
+5. Count cells using a hemocytometer or cell counter.
+
+6. Seed 350 million cells from the CS2 into one CS5 with a total volume of 850 mL.
+   Return to the incubator for 24 h–36 h.
+
+7. Proceed with transfection:
+   • Calculate the amount of each plasmid needed to have a 1:1:1 molar ratio with 2
+     mg total DNA per CS5
+
+     Example calculations:
+     - RepCap (7,265 bp): 582.1 μg = 582.1 μL (at 1.00 μg/μL)
+     - pHelper (11,854 bp): 949.8 μg = 949.8 μL (at 1.00 μg/μL)
+     - Transfer Plasmid (5,842 bp): 468.1 μg = 468.1 μL (at 1.00 μg/μL)
+
+   • Aliquot 100 mL of OptiMEM into a sterile 250 mL bottle.
+
+   • Aliquot 774 mL of DMEM + 2% HI-FBS into a sterile 1 L bottle.
+
+   • Add each plasmid DNA into the bottle containing the Opti-MEM. Mix well.
+
+   • Add 4 mL of PEI (1:2 μg DNA to μg PEI ratio). Shake the bottle up/down
+     vigorously for 30 sec (it's okay to make bubbles).
+
+   • Incubate at RT for 15 min. Note that longer incubation times can reduce
+     transfection efficiency.
+
+   • Add the OptiMEM + DNA + PEI solution to the bottle containing 774 mL of DMEM
+     + 2% HI-FBS. Mix well.
+
+8. Take the CS5 out of the incubator and pour the media into a waste container.
+
+9. Carefully add the OptiMEM + DNA + PEI solution to the CS5. Make sure that all
+   five layers are covered with media. 293T cells are delicate and detach very
+   easily - media should always be added away from the cells (not poured on them)
+   and can be adjusted by carefully tilting the CS5 back and forth.
+
+   💡 PRO TIP: To help distribute the media amongst the five layers, tilt the CS5
+   such that the media goes toward the cap. If the media touches the cap, replace
+   it with a new one before putting the CS5 in the incubator.
+
+10. Return the CS5 to the incubator for 96 h. This incubation time can be adjusted
+    depending upon the serotype. Often AAV2 are harvested at 48–72 h, while other
+    serotypes are harvested at 96–120 h.
+
+11. Optional: perform a media exchange at ~18 h post transfection. Changing the
+    media can reduce toxicity associated with PEI and allow for the addition of
+    other components, such as sorbitol and sodium bicarbonate, sorbitol has been
+    found to increase viral titers by up to 1.8-fold. See the recipe for D1 + 0.1 M
+    sorbitol above.
+    • Carefully pour off the media into a waste container.
+    • Gently pour new media into the CS5.
+    • Distribute the media evenly between the layers.
+    • Return the CS5 to the incubator for ~72 h.
+
+12. Harvest cells and media by tapping the sides of the CS5. Cells should detach
+    easily.
+
+13. Transfer cells and media into 500 mL conical vessels.
+
+14. Rinse CS5 once with 100 mL of PBS and add to the harvested media and cells.
+
+15. Centrifuge at 3900 rpm for 20 min at 4 °C to pellet the cells.
+
+16. Keep the cell pellet on ice and transfer supernatant to a sterile 500 mL
+    bottle.
+
+17. Process the supernatant as follows:
+    • Filter through a 0.45 μm PES membrane.
+    • Add 40% PEG such that the final PEG concentration is 8%.
+    • Add 25 mL of PEG solution to each 100 mL of supernatant. Split into 2 x 500
+      mL sterile bottles as needed.
+    • Add stir bar and stir slowly at 4 °C for 1 h, then keep at 4 °C for 3 h
+      without stirring to allow full precipitation. Precipitation of the viruses
+      can proceed overnight at 4 °C if needed.
+    • Transfer the entire sample to 3 x 500 mL conical bottles and centrifuge at
+      3900 rpm for 15 min at 4 °C.
+    • Discard the supernatant and resuspend the pellets in a total of 5 mL of cell
+      lysis buffer (recipe above). Pipet back and forth to resuspend each pellet
+      completely.
+    • Combine resuspended pellets and keep on ice.
+
+18. Process the cell pellet from above as follows:
+    • Resuspend and lyse the cells by adding a total of 12.5 mL of cell lysis
+      buffer. Pipet back and forth to resuspend each pellet completely.
+    • Combine all resuspended cell pellets and sonicate 5 x 1 sec pulses with at
+      least 5 minutes on ice between each pulse, 50% amplitude. Return to ice
+      between each round of sonication to avoid overheating of the sample. Mix
+      well between rounds of sonication.
+    • Sonicate until no live cells can be seen when stained with Trypan Blue.
+    • Pellet cell debris by centrifugation at 3900 rpm for 10 min at 4 °C.
+    • Transfer the cleared lysate to the tube containing the resuspended virus from
+      step 17 above.
+
+19. Add 50 units of benzonase per mL of viral suspension (you should have ~18 mL
+    from steps 17 & 18). Benzonase is an endonuclease that will degrade any
+    residual DNA carried over from the packaging process.
+
+20. Incubate at 37 °C for 45 min.
+
+21. You are now ready to purify your prep. The clarified supernatant can be kept
+    overnight at 4 °C before proceeding with the purification protocol.
+
+
+SAMPLE DATA
+================================================================================
+
+HEK293T cells at various confluencies. This AAV production protocol should be
+started with cells that are ~80% confluent (center panel). These cells were
+imaged with a 10X objective and were split 2 days before these images were taken.
+
+Last reviewed on: October 10, 2023

biomni/tool/protocols/addgene/Addgene_ AAV Purification.txt

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+PROTOCOL: AAV Purification
+SOURCE: https://www.addgene.org/protocols/aav-purification-iodixanol-gradient-ultracentrifugation/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+This protocol can be used to purify AAV of any serotype.
+
+In medicine, iodixanol can be used as an intravenous isomolar contrast agent. In
+research, iodixanol is used as an isomolar density gradient medium suitable for
+virus purification and isolation of cells, organelles, lipoproteins, and
+macromolecules. Importantly, iodixanol is inert and non-toxic to mammalian cells.
+Therefore, and unlike other density gradient media, it is reportedly not
+necessary to remove iodixanol prior to use of your purified virus. Nonetheless,
+we recommend performing a buffer exchange before using the purified AAV in vivo.
+
+The iodixanol gradient in this protocol is composed of steps that separate out
+contaminants from an impure AAV preparation. The 15% iodixanol step has 1M NaCl to
+destabilize ionic interactions between macromolecules. The 40% and 25% steps are
+used to remove contaminants with lower densities, including empty capsids. The 60%
+step serves as a cushion for genome-containing virions. Phenol red is added to
+clearly distinguish the steps. This method will help enrich the prep with full
+(genome containing) particles. On average, iodixanol-purified AAV preparations
+contain ~20% empty particles.
+
+60% iodixanol solution is available under the name OptiPrep from Sigma-Aldrich.
+
+
+WORKFLOW TIMELINE
+================================================================================
+
+Day 1: Purify
+
+Day 2: Buffer exchange and concentration
+
+Note: Both steps could be completed in one (long) day.
+
+
+EQUIPMENT
+================================================================================
+
+• Ultracentrifuge
+• T70i rotor or equivalent
+
+
+REAGENTS
+================================================================================
+
+• Opti-Prep (60% Iodixanol)
+• QuickSeal tubes
+• QuickSeal tube spacers
+• 16 ga needle
+• 18 ga needle
+• 10 mL syringe
+• 18 ga blunt edge needles, Hamilton
+• 1X PBS pH 7.4
+• 1X PBS-MK buffer
+• 100X Poloxamer 188
+• NaCl
+• MgCl₂
+• KCl
+• Centrifugal filter units (MWCO 100 kDa)
+
+
+REAGENT PREPARATION
+================================================================================
+
+1 M NaCl/PBS-MK buffer:
+
+Dissolve 5.84 g of NaCl, 26.3 mg of MgCl₂ and 14.91 mg of KCl in 1× PBS in a
+final volume of 100 mL. Sterilize by passing through a 0.22 μm filter and store
+at 4 °C.
+
+1X PBS-MK buffer:
+
+Dissolve 26.3 mg of MgCl₂, and 14.91 mg of KCl in 1× PBS in a final volume of
+100 mL. Sterilize by passing through a 0.22 μm filter and store at 4 °C.
+
+0.1% Poloxamer 188 in PBS (A):
+
+Add 500 µL of 10% Poloxamer 188 to 49.5 mL PBS
+
+0.01% Poloxamer 188 in PBS (B):
+
+Add 5 mL of Buffer A to 45 mL PBS
+
+0.001% Poloxamer 188 in PBS + 200 mM NaCl (C) (formulation buffer):
+
+Add 5 mL of Buffer B and 2 mL of 5 M NaCl to 43 mL PBS
+
+
+PROCEDURE
+================================================================================
+
+Preparation and loading of the iodixanol gradient:
+
+1. Prepare the gradient steps:
+   • 15% iodixanol step: mix 4.5 mL of 60% iodixanol and 13.5 mL of 1 M
+     NaCl/PBS-MK buffer
+   • 25% iodixanol step: mix 5 mL of 60% iodixanol and 7 mL of 1X PBS-MK buffer
+     and 30 μL of phenol red
+   • 40% iodixanol step: mix 6.7 mL of 60% iodixanol and 3.3 mL of 1X PBS-MK
+     buffer
+   • 60% iodixanol step: mix 10 mL of 60% iodixanol and 45 μL of phenol red
+
+2. Underlay each solution into a QuickSeal tube in the order below using a 10 mL
+   syringe and a blunt edge 18 ga Hamilton needle, taking care to avoid bubbles:
+   • 8 mL of 15% iodixanol step
+   • 5 mL of 25% iodixanol step
+   • 5 mL of 40% iodixanol step
+   • 3 mL of 60% iodixanol step
+
+3. Carefully add up to 5 mL of clarified supernatant on top of the gradient. Use
+   1X PBS (or formulation buffer) to top off the tube.
+
+4. Seal the QuickSeal tubes.
+
+5. Centrifuge at 350,000 x g for 90 min in a T70i rotor at 10 °C.
+
+   💡 PRO TIP: If you need more time, you can alternatively centrifuge for 2 h at
+   200,000 x g at 18 °C.
+
+6. Carefully take the QuickSeal tubes out of the rotor and place them in a stable
+   rack.
+
+   **Make sure not to disturb the gradient!**
+
+Collect Fractions
+
+Option #1:
+
+1. Prepare a row of roughly 20 open 1.5 mL microcentrifuge tubes in a rack.
+   These will be used to collect fractions from the gradient. You can collect fewer
+   fractions once you have a good idea when the virus elutes from the gradients.
+
+2. In a biosafety cabinet, carefully puncture the QuickSeal tube at the interface
+   of the 60% and 40% gradient (see Figure 2) with an 18 ga needle.
+
+3. Place the first microcentrifuge tube under the needle's opening to collect the
+   fractions.
+
+4. Puncture the top of the QuickSeal tube with a 16 ga needle and start
+   collecting 0.5 mL to 1 mL fractions per tube.
+
+5. Avoid the proteinaceous material near the 40–25% interface.
+
+   💡 PRO TIP: As soon as the top of the tube is punctured, AAV-containing
+   Iodixanol solution will flow from the needle set at the 40–60% interface.
+   Make sure that the microcentrifuge tubes are well positioned to collect the
+   solution, or you will lose a significant amount of your virus. When the first
+   fraction is collected, move the rack to the next empty tube to collect the next
+   fraction. The flow will be rapid at first and will progressively slow down.
+
+Option #2:
+
+1. Puncture the QuickSeal tube at the bottom using an 18 ga needle.
+
+2. Immediately start collecting 0.5 to 1 mL fractions in microcentrifuge tubes.
+
+3. Repeat for each QuickSeal tube.
+
+4. The first 3 mL collected corresponds to the 60% phase and can be discarded.
+
+5. The fractions obtained from the 40% phase contain the purified AAV.
+
+Option #3:
+
+1. Puncture the QuickSeal tube slightly below the 60–40% interface with an 18 ga
+   needle attached to a 10 mL syringe.
+
+2. The bevel of the needle should be up, facing the 40% step.
+
+3. Collect up to 5 mL per tube, taking care to avoid collecting the
+   proteinaceous material at the 40–25% interface.
+
+4. Repeat for each QuickSeal tube.
+
+5. (Optional) For first time users, it is a good idea to assay each fraction by
+   silver stain or SYPRO Ruby stain to determine purity. Only the cleanest
+   fractions should be kept and pooled for dialysis.
+
+6. Pool clean fractions.
+
+Concentration and buffer exchange:
+
+Stocks of Poloxamer 188:
+• Poloxamer 188, 10% solution
+• A: 0.1% in PBS: 49.5 mL PBS + 500 µl Poloxamer 188
+• B: 0.01% in PBS: 45 mL PBS + 5 ml A
+• C: 0.001% in PBS: 45 mL PBS + 5 ml B + 200 mM NaCl
+
+Procedure:
+
+1. Cover the filter membrane with 15 mL of 0.1% Poloxamer 188 PBS and incubate
+   for 10 min at RT
+
+2. Spin at 3000 rpm for 5 min at 4 °C and discard the flow through.
+
+3. Add 15 mL of 0.01% Poloxamer 188 PBS and incubate for 10 min at RT.
+
+4. Spin at 3000 rpm for 5 min at 4 °C and discard the flow through.
+
+5. Add 15 mL of 0.001% Poloxamer 188 + 200mM NaCl PBS.
+
+6. Spin at 3000 rpm for 5 min at 4 °C and discard the flow through.
+
+7. Add your sample.
+
+8. Spin at 3500 rpm for 5–8 min at 4 °C, discard the flow through.
+
+9. Add more sample and spin 3500 rpm for 2–5 min at 4 °C, discard the flow
+   through.
+
+10. Repeat this step as needed.
+
+    Please note that iodixanol is not easily removed. After each spin, add more
+    formulation buffer and sample and make sure to pipet back and forth a few
+    times to mix the iodixanol that has settled at the bottom of the column.
+
+11. We recommend concentrating to a minimum of 500 µL. If the concentrate volume
+    is less than 500 µL, bring up the volume with formulation buffer.
+
+12. Use a P1000 to the bottom of the filter and pipette up/down and wash off the
+    walls of the filter to recover as much virus as possible.
+
+13. Store at 4 °C for short term (2 weeks), or aliquot and store at -80 °C for
+    long term.
+
+
+SAMPLE DATA
+================================================================================
+
+Iodixanol gradient before ultracentrifugation. The steps are clearly
+distinguishable thanks to the use of phenol red.
+
+Iodixanol gradient after ultracentrifugation. The arrow indicates the 60–40%
+interface. The vertical black line indicates the location of the purified AAV.
+
+Example of an SDS-PAGE gel of 10 consecutive gradient fractions followed by
+silver stain. Note the increased number of contaminants in each fraction. * AAV
+capsid proteins VP1, VP2, VP3; M protein marker.
+
+Last reviewed on: November 7, 2023

biomni/tool/protocols/addgene/Addgene_ AAV Titration by qPCR.txt

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+PROTOCOL: AAV Titration by qPCR
+SOURCE: https://www.addgene.org/protocols/aav-titration-qpcr-using-sybr-green-technology/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+The AAV Titration protocol can be used to determine the number of
+genome-containing particles of an AAV prep using SYBR green technology.
+
+This protocol was validated using an internal reference AAV of known titer,
+100837-AAV1, and by measuring the titer of samples obtained from academic viral
+vector cores. Our titers were similar to those reported by these institutions.
+
+This protocol is for a 96-well plate with 20 μL reaction volume.
+
+Last Update: February 13, 2019
+
+Estimate of time required: ~3 h
+
+
+WORKFLOW TIMELINE
+================================================================================
+
+Plate set-up: 2 h
+
+qPCR run: 1.5 h
+
+Data analysis: 30 min
+
+
+EQUIPMENT
+================================================================================
+
+• qPCR instrument
+• Heating plate
+• Pipettors
+• 1–10 µL single channel pipette
+• 20–200 µL single channel pipette
+• 200–1000 µL single channel pipette
+• 1–10 µL multichannel pipette
+• 2–50 µL multichannel pipette
+• 20–200 µL multichannel pipette
+
+
+REAGENTS
+================================================================================
+
+• Universal SYBR Master Mix 2X
+• Primer pair targeting AAV2 ITR (Aurnhammer et al., 2012)
+  - fwd ITR primer, 5'-GGAACCCCTAGTGATGGAGTT
+  - rev ITR primer, 5'-CGGCCTCAGTGAGCGA
+• ITR-containing plasmid for standard curve
+• RNase-free DNase
+• 10X DNase buffer
+• Nuclease-free water
+• Microcentrifuge tubes
+• 96-well optical plate
+• Pipette tips
+
+
+GENERAL CONSIDERATIONS
+================================================================================
+
+• Always run standards and samples in duplicate at least
+
+• If possible, include an AAV reference sample of known titer. The reference
+  material should have a titer within 1-log of the expected titer of the samples
+  being tested.
+
+• Always include a No Template Control (NTC), i.e master mix + water
+
+• Whenever possible use a multichannel pipet to minimize pipetting error and
+  variability
+
+• Mix samples very well by pipetting back and forth multiple times at each step
+
+
+REAGENT PREPARATION
+================================================================================
+
+Master Mix:
+
+Count the number of samples (n) and prepare master mix for an additional 10
+samples (n+10 – the additional amount will ensure that there is enough master mix
+for all samples). Each sample requires 15 μL of master mix.
+
+💡 PRO TIPS:
+
+• Use a "Universal" SYBR master mix which contains a high-quality DNA polymerase
+  and a blend of dTTP/dUTP to minimize carryover contamination. The master mix
+  should also contain an internal passive reference (typically ROX dye), to
+  normalize non-PCR–related fluorescence fluctuations and to minimize well-to-well
+  variability that result from a variety of causes, such as pipetting error and
+  sample evaporation.
+
+• Make the master mix after all the samples have been added to the qPCR plate.
+  Start by adding water, then SYBR master mix, then the forward and reverse
+  primers. Vortex briefly, immediately before use.
+
+• Use a reservoir and a multichannel pipette to dispense the master mix into the
+  wells.
+
+Master Mix Recipe:
+
+Reagent                    | Amount for ONE Reaction | Amount for 100 reactions
+---------------------------|------------------------|--------------------------
+Universal SYBR Master Mix  | 10 μL                  | 1,000 μL
+2X                         |                        |
+100 μM Forward Primer      | 0.15 μL                | 15 μL
+100 μM Reverse Primer      | 0.15 μL                | 15 μL
+Nuclease Free Water        | 4.7 μL                 | 470 μL
+
+
+PROCEDURE
+================================================================================
+
+1. Prepare a plasmid stock of 2 x 10⁹ molecules/μL to generate a standard curve:
+
+   One option is to use plasmid #59462 from Addgene. The values highlighted below
+   in red were calculated using this plasmid, but will change if you use a
+   different plasmid.
+
+   Sample Calculation:
+   • Size in bp of Addgene plasmid #59462: 6208 bp
+   • Concentration of Addgene plasmid #59462: 1.07 μg/μL
+   • Molecular Weight: 6208 bp × 650 dalton/bp = 4.03 × 10⁶ g/mole
+   • Moles/μl: 1.07 μg/μL × 1 g/10⁶ μg × 1 / 4.03 × 10⁶ g/mole = 2.65 × 10⁻¹³
+     moles/μL
+   • Molecules/μl: 2.65 × 10⁻¹³ moles/μL × 6.022145 × 10²³ molecules/mole = 1.59
+     × 10¹¹ molecules/μL
+   • To obtain a solution at 2 × 10⁹ molecules/μL: 1.59 × 10¹¹ / 2 × 10⁹ = 79.8X
+     dilution
+   • Therefore we need to dilute 1.25 μL stock into 98.74 μL H₂O
+
+   💡 PRO TIP: Once a validated standard curve is obtained, make a small aliquot
+   of each standard (enough for 1 or 2 plates) and store at -20 °C. Once a
+   standard is thawed do not freeze it again but store at 4 °C and use within 1
+   week.
+
+   Keep track of the Ct value for each standard over time. They should remain
+   within 0.5 Ct of their initial value. If the Ct value of the standard starts
+   to drift, it's time to make a new one.
+
+   When developing the assay multiple plasmids containing ITR were tested.
+   Plasmid #59462 is one plasmid that gave reliable and consistent results. Use
+   the recommended plasmid, or test multiple plasmids to find a suitable one.
+
+   Some labs have reported better results when the plasmid is linearized.
+
+2. Treat the purified AAV samples with DNase I to eliminate any contaminating
+   plasmid DNA carried over from the production process (DNase does not penetrate
+   the virion).
+
+   • 5 μL sample + 39 μL H₂O + 5 μL 10X DNase buffer + 1 μL DNase
+   • Gently mix sample (do not vortex)
+   • Incubate 30 min at 37 °C
+   • Transfer to ice
+
+   ** Critical: do NOT treat your plasmid standard with DNase **
+
+3. Make 6 serial dilutions, in duplicate, of your standard curve plasmid (2 ×
+   10⁹ stock made in step #1):
+
+   Volume of 2 × 10⁹ stock | Volume of nuclease-free | Molecules per μL
+   or previous dilution   | water (uL)              |
+   (μL)                    |                         |
+   ------------------------|-------------------------|------------------
+   10                      | 90                      | 2 × 10⁸
+   10 of 2 × 10⁸ dilution  | 90                      | 2 × 10⁷
+   10 of 2 × 10⁷ dilution  | 90                      | 2 × 10⁶
+   10 of 2 × 10⁶ dilution  | 90                      | 2 × 10⁵
+   10 of 2 × 10⁵ dilution  | 90                      | 2 × 10⁴
+   10 of 2 × 10⁴ dilution  | 90                      | 2 × 10³
+
+   💡 PRO TIP: To help stabilize the standards add carrier DNA to a final
+   concentration of 4 ug/mL to each standard dilution.
+
+4. Dilute DNase-treated and AAV reference samples according to the dilution
+   scheme in the table below:
+
+   Dilution Series | Volume of sample | Volume of nuclease | Dilution | Total
+                   | (uL)             | free water (uL)    | factor   | dilution
+   ----------------|------------------|--------------------|----------|----------
+   Dilution 1      | 5 uL AAV stock   | 45 uL              | 10X      | 10X
+   (DNase step)    |                  |                    |          |
+   Dilution 2      | 5 uL Dil. 1     | 95 uL              | 20X      | 200X
+   Dilution 3      | 20 uL Dil. 2    | 80 uL              | 5X       | 1000X
+   Dilution 4      | 20 uL Dil. 3    | 80 uL              | 5X       | 5000X
+   Dilution 5      | 20 uL Dil. 4    | 80 uL              | 5X       | 25000X
+   Dilution 6      | 20 uL Dil. 5    | 80 uL              | 5X       | 125000X
+   Dilution 7      | 20 uL Dil. 6    | 80 uL              | 5X       | 625000X
+   Dilution 8      | 20 uL Dil. 7    | 80 uL              | 5X       | 3125000X
+
+   Dilutions highlighted in green are the ones loaded onto the qPCR plate for
+   most samples.
+
+   • If sample is expected to have a titer <1 × 10¹² GC/mL, use dilutions 3–6
+   • If sample is expected to have a titer >3 × 10¹³ GC/mL, use dilutions 5–8
+
+   Note: at Addgene, samples typically range from 1 × 10¹² GC/mL to >2 × 10¹³
+   GC/mL and we use an internal reference virus that is 1 × 10¹³ GC/mL. We
+   recommend always using a reference within 1-log of the expected titer.
+
+   💡 PRO TIPS:
+   • The quality of the sample dilution series is critical. Make sure to pipet
+     each dilution up and down at least 10 times, and use at least half of the
+     final volume (mix with >50 uL if your well contains 100 uL)
+   • Use a multichannel pipette to load the standards and samples onto the qPCR
+     plate
+
+5. Set up and load the 96-well plate:
+   • Load 5 μL of each standard in duplicate
+   • Load 5 μL of each sample in duplicate. Do not forget to include a no
+     template control (NTC = master mix + water).
+   • Add 15 μL of Master Mix per well and mix well by pipetting back and forth
+     at least 5 times.
+   • Seal plate with transparent film.
+   • Centrifuge at 3,000 rpm for 2 min to bring the sample to the bottom of the
+     tube.
+
+6. Run the following protocol in your qPCR instrument using SYBR detection:
+
+   98 °C 3 min / 98 °C 15 sec / 58 °C 30 sec / read plate/ repeat 39x from step
+   3 / melt curve
+
+7. Perform data analysis using the instrument's software. Determine the physical
+   titer of samples (viral genomes (vg)/mL) based on the standard curve and the
+   sample dilutions.
+
+   💡 PRO TIP: Make sure that the qPCR is valid by checking to the following:
+   • Standard curve: R² (coefficient of correlation) ~ 1.0, E (efficiency of PCR)
+     ~100% (90%–110% range is acceptable)
+   • Baseline removal: all samples will have some small amount of background
+     signal that is most evident during initial PCR cycles. This background
+     signal must be removed to accurately determine differences between samples.
+   • Melt curve analysis: a single peak should be seen. The presence of a second
+     peak at a temperature of ~70–75 °C usually indicates the presence of primer
+     dimers which can increase background signal and alter the Ct values of your
+     samples.
+   • Quality of your standard curve: you should observe differences in Ct values
+     that make sense for your dilutions (~3.3 difference Ct for a 10-fold dilution
+     is appropriate).
+   • Quality of duplicates: Exclude duplicates from analyses if there is more than
+     a 0.5 Ct difference between them.
+
+
+SAMPLE DATA
+================================================================================
+
+Example of a valid 8-point standard curve.
+
+Example of the amplification plots obtained from an AAV sample. Each curve
+represents a dilution.
+
+
+REFERENCES
+================================================================================
+
+Aurnhammer C, Haase M, Muether N, Hausl M, Rauschhuber C, Huber I, Nitschko H,
+Busch U, Sing A, Ehrhardt A, Baiker A. Universal real-time PCR for the detection
+and quantification of adeno-associated virus serotype 2-derived inverted
+terminal repeat sequences. Hum Gene Ther Methods. 2012 Feb;23(1):18-28. PMID:
+22428977

biomni/tool/protocols/addgene/Addgene_ Affinity Purification of Recombinant Antibodies with Protein A or Protein G.txt

@@ -0,0 +1,310 @@
+PROTOCOL: Affinity Purification of Recombinant Antibodies with Protein A or Protein G
+SOURCE: https://www.addgene.org/protocols/affinity-purification-recombinant-antibodies/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+This protocol describes how to affinity purify recombinant antibodies from cell
+culture supernatant using Protein A or Protein G columns.
+
+Last Update: February 7, 2022
+
+
+SAFETY WARNINGS
+================================================================================
+
+Sodium azide is toxic. When handling, use proper personal protective equipment
+including laboratory coat, laboratory goggles, and gloves. Sodium azide containing
+waste should be properly disposed of following your institution's hazardous waste
+procedures.
+
+
+WORKFLOW TIMELINE
+================================================================================
+
+Day 1: Purify antibody
+
+Day 2 or later: Buffer exchange
+
+
+EQUIPMENT
+================================================================================
+
+• Class II, Type A2 Biological Safety Cabinet
+• 4 °C Refrigerator
+• Pipette controller
+• Benchtop centrifuge compatible with 50 mL conical tubes
+• NanoDrop spectrophotometer
+• 37 °C, 5% CO₂ incubator with shaking platform set to 120 rpm
+• 37 °C bead bath
+• Clamp stand and clamps
+• Autoclave
+• 0.1–1 mL single channel pipette
+• 0.5–10 µL single channel pipette
+• 20–200 µL single channel pipette
+• 2–20 µL single channel pipette
+
+
+REAGENTS
+================================================================================
+
+• Aspirating pipette, VWR 414004-265
+• 5 mL pipette, VWR 89130-896
+• 10 mL pipette, VWR 89130-898
+• 25 mL pipette, VWR 89130-900
+• 50 mL pipette, VWR 89130-902
+• 50 mL conical tubes, VWR 89039-656
+• Microcentrifuge tubes, VWR 89126-724
+• Filtering system 0.2 µm PES, VWR 431098
+• Sodium phosphate monobasic monohydrate, Millipore Sigma 71507-250G
+• Sodium phosphate dibasic, Millipore Sigma S7907-500G
+• 1 M Trizma hydrochloride solution pH 9.0, Millipore Sigma T2819-1L
+• 250 mL sterile bottle, VWR 430281
+• 0.45 µm PES complete filtration unit, 500 mL, rapid-flow, VWR 73520-984
+• Syringes, 30 mL, VWR BD302832
+• Filter, 0.2 µm (luer-lock), VWR 431229
+• IgG Elution Buffer, Thermo Fisher 21028
+• Protein G GraviTrap columns, Cytiva 28-9852-55 or rProtein A GraviTrap
+  columns, Cytiva 28-9852-54
+• LabMate PD10 Buffer Reservoir, Millipre Sigma GE18-3216-03
+• 5% Sodium Azide, 500 mL, VWR 7144.8-16
+• PBS, 1X pH 7.4, VWR 45000-446
+• Benzamide, Millipore Sigma 12072
+• Antipain, Millipore Sigma 10791
+• Leupeptin, Millipore Sigma L2884
+• Aprotinin, Millipore Sigma A6279
+• Zeba Spin Desalting Columns, 7 kDa MWCO, 10 mL, Thermo Fisher 89893
+• Amicon Ultra-15 30 kDa MWCO column, Millipore Sigma UFC903024
+• 50 mL LoBind tubes, VWR 76289-498
+
+
+BEFORE STARTING
+================================================================================
+
+• Wipe down all pipettes and equipment with 10% bleach prior to use.
+
+• Warm the affinity column, binding buffer, and elution buffer to room
+  temperature (RT) before use.
+
+
+REAGENT PREPARATION
+================================================================================
+
+1 M sodium phosphate monobasic monohydrate (NaH₂PO₄∙H₂O), pH 7.0:
+
+• 138 g NaH₂PO₄∙H₂O
+• 1 L deionized water
+• Adjust pH to 7.0
+• Autoclave or filter sterilize
+
+1 M sodium phosphate dibasic (Na₂HPO₄), pH 7.0:
+
+• 142 g of Na₂HPO₄∙H₂O
+• 1 L deionized water
+• Adjust pH to 7.0
+• Autoclave or filter sterilize
+
+1 M of sodium phosphate buffer, pH 7.0:
+
+• 390 mL of sterile sodium phosphate monobasic monohydrate (NaH₂PO₄∙H₂O)
+• 610 mL of sterile sodium phosphate dibasic (Na₂HPO₄)
+• Store up to 1 month at 4 °C
+• Adjust pH to 7.0
+• Autoclave or filter sterilize
+
+Protein A/G binding buffer (20 mM sodium phosphate buffer, pH 7.0):
+
+• 20 mL 1M sodium phosphate buffer
+• 980 mL of deionized water
+• Adjust pH to 7.0
+• Autoclave or filter sterilize
+
+1000X protease inhibitor cocktail:
+
+• 25 mg leupeptin
+• 50 mg antipain
+• 250 mg benzamidine
+• 25 mL of 2 mg/mL aprotinin
+• Mix well and sterilize through a 0.2 µm PES filter.
+• Aliquot and freeze upright at -20 °C
+
+
+PROCEDURE
+================================================================================
+
+Section 1: Affinity chromatography
+
+1. Equilibrate filtered tissue culture supernatant containing the recombinant
+   antibody to room temperature (see Transfection for Recombinant Antibody
+   protocol).
+
+   💡 PRO TIP: Add protease inhibitors to 1X if this has not already been done.
+   We typically start with about 250 mL of supernatant and add 250 µL of 100X
+   protease inhibitor cocktail.
+
+2. Add 1 part Protein A/G binding buffer to 1 part tissue culture supernatant
+   and mix well.
+
+3. Uncap the Protein G Gravitrap or rProtein A Gravitrap columns and gently pour
+   off the ethanol storage buffer.
+
+4. Tightly attach LabMate PD10 Buffer Reservoirs to the top of the Gravitrap
+   column.
+
+   💡 PRO TIP: Gravitrap columns have a finite binding capacity. If your sample
+   exceeds the capacity, divide the sample among multiple columns. For
+   information about the total binding capacity for your columns, refer to the
+   manufacturer's instructions.
+
+5. Attach the Gravitrap column to a clamp on a clamp stand.
+
+6. Use scissors to cut the bottom of the Gravitrap column at the indentation in
+   the tip.
+
+7. Equilibrate the column with 10 mL of RT Protein A/G binding buffer.
+
+8. Collect the flow-through in a 50 mL conical tube and dispose after use.
+
+9. Carefully pour the filtered and diluted cell culture supernatant into the
+   Gravitrap column.
+
+10. Collect the flow through in a 250 mL bottle.
+
+11. Repeat steps 9–10 such that the supernatant passes through the column a total
+    of 2x.
+
+    💡 PRO TIP: This will ensure depletion of the recombinant antibody from the
+    supernatant.
+
+12. Wash the column with 25 mL of Protein A/G binding buffer 2x.
+
+13. Add 50 µL of 1 M Trizma hydrochloride pH 9.0 to 10 microcentrifuge tubes.
+
+14. Cap the Gravitrap columns at the bottom.
+
+15. Add 5 mL of Pierce IgG Elution Buffer to the capped Gravitrap column and
+    incubate for 10 min at RT.
+
+16. Uncap the column and collect 500 µL fractions into the tubes containing 50 µL
+    1 M Trizma hydrochloride pH 9.0.
+
+17. Cap the tubes and vortex for 5 s to mix.
+
+18. Determine the protein concentration of each fraction on the NanoDrop
+    Spectrophotometer using the A280 IgG setting.
+
+19. Combine all eluates with measurable protein.
+
+20. Determine the protein concentration of the pooled sample on the NanoDrop
+    Spectrophotometer using the A280 IgG setting.
+
+21. If the concentration of the pooled sample is above 1.0 mg/mL proceed to Option
+    1 with a buffer exchange using a Zeba Spin 7 kDa MWCO desalting column.
+
+22. If the concentration of the pooled sample is below 1.0 mg/mL proceed to
+    Option 2 with a buffer exchange/concentration using an Amicon 30 kDa MWCO buffer
+    exchange column.
+
+23. (Optional) Regenerate the column by washing with 25 mL of Protein A/G binding
+    buffer and store in 20% ethanol at 4 °C.
+
+    💡 PRO TIP: Columns may be re-used up to 5x when purifying the same
+    recombinant antibody.
+
+Section 2: Buffer exchange
+
+Choose Option 1 or Option 2 based on the concentration of the pooled sample above.
+
+Option 1: Buffer exchange using a desalting column
+
+The 10 mL Zeba Spin Desalting Columns can accommodate up to 4 mL of recombinant
+antibody sample.
+
+💡 PRO TIP: If the volume of the sample is greater than 4 mL, then divide it into
+2 columns.
+
+1. Twist off the 10 mL Zeba Spin Desalting Column's bottom closure and loosen cap.
+
+2. Place column in a 50 mL conical collection tube.
+
+3. Centrifuge column at 1000 x g for 2 min to remove storage solution.
+
+4. Place a mark on the side of the column where the compacted resin is slanted
+   upward. Place column in centrifuge with the mark facing outward in all
+   subsequent centrifugation step
+
+   💡 PRO TIP: Resin will appear compacted after centrifugation.
+
+5. Discard the flow through into a waste container.
+
+6. Add 5 mL PBS to the column.
+
+7. Centrifuge at 1000 x g for 2 min to remove PBS.
+
+8. Repeat Steps 6–7 3x, discarding buffer from the collection tube each time.
+
+9. Place column in a new 50 mL LoBind collection tube.
+
+10. Slowly apply the sample to the center of the compact resin bed.
+
+    💡 PRO TIP: For sample volumes <1.5 mL top off the sample with 0.2 mL PBS to
+    aid protein recovery.
+
+11. Allow the resin bed to fully absorb the sample.
+
+12. Centrifuge at 1000 x g for 2 min to collect the sample.
+
+13. Discard column after use.
+
+14. Determine antibody concentration on the NanoDrop Spectrophotometer.
+
+15. Dilute antibody to 1 mg/mL with PBS if needed.
+
+16. For long term storage, add sterile sodium azide to 1 mM.
+
+Option 2: Buffer exchange and concentration with an Amicon column
+
+1. Apply 15 mL of PBS to the reservoir of a Amicon Ultra-4 30 kDa MWCO column.
+
+2. Incubate for 10 min at RT.
+
+3. Centrifuge at 3100 x g for 8 min.
+
+4. Discard the flow through into a waste container.
+
+5. Add the recombinant antibody sample to the reservoir of the column.
+
+6. Fill the remaining space in the column with PBS and pipette several times to
+   mix.
+
+7. Centrifuge at 3100 x g for 8 min.
+
+8. Discard the flow through into a waste container.
+
+9. Repeat steps 5-8 at least 4 additional times to ensure a full buffer exchange.
+
+10. Discard the flow through into a waste container.
+
+11. Fill the remaining space in the column with PBS and pipette several times to
+    mix.
+
+12. Centrifuge at 3100 x g for 8 min.
+
+13. Remove a small aliquot of your sample from the filter reservoir to check the
+    concentration on the NanoDrop Spectrophotometer to see if the sample has
+    reached the desired concentration.
+
+14. If the sample concentration is still too low, repeat the centrifugation until
+    the volume of the sample is reduced enough to reach the desired concentration.
+
+    💡 PRO TIP: Periodically check the concentration on the NanoDrop
+    Spectrophotometer to see if the sample has reached the desired concentration.
+
+15. Gently transfer the sample from the reservoir to a LoBind tube.
+
+    💡 PRO TIP: Do not scrape the filter with the pipette tip.
+
+16. Determine the concentration of the sample on the NanoDrop Spectrophotometer.
+
+17. For long term storage, add sterile sodium azide to 1 mM.

biomni/tool/protocols/addgene/Addgene_ Antibody Validation Using the Indirect ELISA Method.txt

@@ -0,0 +1,280 @@
+PROTOCOL: Antibody Validation Using the Indirect ELISA Method
+SOURCE: https://www.addgene.org/protocols/ab-validation-indirect-elisa/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+This protocol describes how to run an indirect Enzyme-Linked ImmunoSorbent Assay
+(ELISA) against a purified antigen to validate an antibody. This protocol will
+describe how to create a standard curve using known concentrations of the antigen
+and how to probe with an antibody against the target to show a dose response.
+
+
+GENERAL CONSIDERATIONS
+================================================================================
+
+The ELISA can be completed in a single day or broken up over several days by
+incubating certain steps at 4 °C instead of room temperature or 37 °C. The
+protocol notes when there are different incubation options available.
+
+Caution: Use the Stop Solution in a well ventilated area and avoid breathing in
+the vapors.
+
+
+WORKFLOW TIMELINE
+================================================================================
+
+Day 1: Antigen Coating
+
+Day 2: Blocking
+
+Day 3: Primary antibody incubation
+
+Day 4: Secondary antibody incubation and plate read
+
+
+EQUIPMENT
+================================================================================
+
+• Spectrophotometer compatible with 96-well plates
+• 1–10 µL single channel pipette
+• 2–20 µL single channel pipette
+• 20–200 µL single channel pipette
+• 200–1000 µL single channel pipette
+• 20–200 µL multichannel pipette
+• Multichannel pipette reagent reservoirs
+• Pipette controller
+• Microcentrifuge
+• Microplate shaker
+• ELISA plate washer (optional)
+• Scale
+• Microsoft Excel or similar software
+
+
+REAGENTS
+================================================================================
+
+• Purified antigen
+• Purified recombinant antibody
+• Isotype control antibody
+• HRP-conjugated isotype-specific secondary antibody
+• ELISA 96-well microplate, Corning 9018
+• Bovine serum albumin (BSA), VWR 10273-264
+• 50 mL conical tubes, VWR 89039-656
+• 96-well polyester (clear) microplate seal, Thermo Scientific 5701
+• Microcentrifuge tubes, Neptune 3745.X
+• Pipette tips, 1000 µL, VWR 76322-154
+• Pipette tips, 10 µL, VWR 76322-132
+• Pipette tips, 200 µL, VWR 76322-150
+• Pipette tips, 20 µL, VWR 76322-134
+• Pipettes, 10 mL, VWR 89130-898
+• 1 L polystyrene bottle, Corning 430518
+• PBS, 1X pH 7.4, VWR 45000-446
+• TMB (3,3',5,5'-tetramethyl-[1,1'-biphenyl]-4,4'-diamine) substrate kit, Pierce
+  34021
+• Tween-20, VWR 0777-1L
+• Foil
+
+
+BEFORE STARTING
+================================================================================
+
+Warm reagents to room temperature.
+
+
+PROCEDURE
+================================================================================
+
+Section 1: Prepare the Antigen Standard and coat the plate
+
+1. Dilute purified antigen to 20 ng/µL in a microfuge tube and vortex.
+
+   💡 PRO TIP: You may need to try a few different concentrations of antigen to
+   determine the ideal concentration.
+
+2. Prepare serial dilutions of the purified antigen as follows:
+   • 2 ng/µL: Add 100 µL of 20 ng/µL stock into 900 µL PBS in a microfuge tube
+     and vortex.
+   • 1 ng/µL: Add 450 µL of 2 ng/µL stock into 450 uL PBS in a microfuge tube and
+     vortex.
+   • 0.5 ng/µL: Add 450 µL of 1 ng/µL stock into 450 µL PBS in a microfuge tube
+     and vortex.
+   • 0.25 ng/µL: Add 450 µL of 0.5 ng/µL stock into 450 µL PBS in a microfuge tube
+     and vortex.
+   • 0.125 ng/µL: Add 450 µL of 0.25 ng/µL stock into 450 µL PBS in a microfuge
+     tube and vortex.
+   • 0 ng/µL: 450 µL PBS only.
+
+3. Pipette 50 µL into the first three wells of each row, using a different
+   standard solution for each row of a 96-well ELISA microplate.
+
+4. Cover the plate with a 96-well plate seal and incubate at 37 °C for 30 min,
+   or overnight at 4 °C.
+
+Section 2: Block the plate
+
+1. Prepare the wash buffer (0.05% Tween-20 in PBS) as follows:
+   • Pipette 0.5 mL of Tween-20 to into 999.5 mL 1X PBS
+   • Cap the bottle and invert several times to mix.
+
+2. Carefully remove the plate seal from the 96-well plate and set aside.
+
+3. Aspirate the wells and use a multichannel pipette to add 200 µL wash buffer to
+   each well. Washing can be done by hand but if available, use an automated ELISA
+   plate washer.
+
+4. Seal the plate.
+
+5. Place the plate on a microplate shaker set at 400 rpm and shake for 1 min.
+
+6. Repeat steps 3–5 twice for a total of three washes.
+
+7. After the final wash, aspirate the wash buffer from the wells.
+
+8. Prepare the blocking buffer (1% BSA in PBS) as follows:
+   • Add 250 mg of BSA to 25 mL of PBS.
+   • Cap the tube and invert several times to mix.
+
+9. Using a multichannel pipette, add 200 µL of blocking buffer to each well.
+
+10. Cover the plate with a plate seal and incubate on a microplate shaker set to
+    400 rpm for 2 h at room temperature or overnight at 4 °C.
+
+Section 3: Primary antibody incubation
+
+1. Carefully remove the plate seal from the 96-well plate and set aside.
+
+2. Aspirate the wells and use a multichannel pipette to add 200 µL wash buffer to
+   each well. Washing can be done by hand but if available, use an automated ELISA
+   plate washer.
+
+3. Seal the plate.
+
+4. Place the plate on a microplate shaker set at 400 rpm and shake for 1 min.
+
+5. Repeat steps 2–4 twice for a total of three washes.
+
+6. After the final wash, aspirate the wash buffer from the wells.
+
+7. Prepare antibody dilution buffer (0.1% BSA in PBS) as follows:
+   • Dilute 50 mg BSA into 50 mL PBS.
+   • Cap the tube and invert several times to mix.
+
+8. Dilute the primary antibody and an isotype control antibody to the desired
+   concentrations in the antibody dilution buffer.
+
+   💡 PRO TIP: The ideal antibody concentration will vary and needs to be
+   empirically determined. We suggest starting with three concentrations between
+   1–10 µg/mL.
+
+9. Cap and invert the tubes several times to mix.
+
+10. Using a multichannel pipette, add 100 µL of primary antibody solution to the
+    appropriate wells.
+
+11. Cover the plate with a plate seal and incubate on a microplate shaker set to
+    400 rpm for 2 h at room temperature or overnight at 4 °C.
+
+Section 4: Secondary antibody incubation
+
+1. Carefully remove the plate seal from the 96-well plate and set aside.
+
+2. Aspirate the wells and use a multichannel pipette to add 200 µL wash buffer to
+   each well. Washing can be done by hand but if available, use an automated ELISA
+   plate washer.
+
+3. Seal the plate.
+
+4. Place the plate on a microplate shaker set at 400 rpm and shake for 1 min.
+
+5. Repeat steps 2–4 twice for a total of three washes.
+
+6. After the final wash, aspirate the wash buffer from the wells.
+
+7. Dilute the HRP-conjugated isotype-specific secondary antibody to the desired
+   concentrations in the antibody dilution buffer.
+
+8. Cap and invert the tube several times to mix.
+
+9. Using a multichannel pipette, add 100 µL of secondary antibody to each well.
+
+10. Cover the plate with a plate seal and incubate on a microplate shaker set to
+    400 rpm for 2 h at room temperature or overnight at 4 °C.
+
+Section 5: TMB reaction
+
+1. Carefully remove the plate seal from the 96-well plate and set aside.
+
+2. Aspirate the wells and use a multichannel pipette to add 200 µL wash buffer to
+   each well. Washing can be done by hand but if available, use an automated ELISA
+   plate washer.
+
+3. Seal the plate.
+
+4. Place the plate on a microplate shaker set at 400 rpm and shake for 1 min.
+
+5. Repeat steps 2–4 twice for a total of three washes.
+
+6. After the final wash, aspirate the wash buffer from the wells.
+
+7. Immediately before use, mix 5 mL TMB Solution and 5 mL Peroxide Solution from
+   the TMB substrate kit.
+
+8. Cap the tube and invert several times to mix.
+
+9. Using a multichannel pipette, add 100 µL of the TMB/Peroxide mix to each well.
+
+10. Cover the plate with a plate seal and wrap in foil.
+
+11. Incubate plate at room temperature on a microplate shaker set at 400 rpm for
+    15–30 min.
+
+    💡 PRO TIP: Check the plate periodically to determine when the desired color
+    change has occurred. The color will change from yellow to blue and generally
+    takes between 15 to 30 min. High concentrations of HRP yield a greenish
+    solution. Stop the reaction before any wells display a green product. A
+    precipitated product indicates the presence of too much HRP and the need to
+    optimize experimental conditions.
+
+12. When the desired color change has occurred, gently remove the plate seal.
+
+13. Using a multichannel pipette, add 100 µL of TMB Stop Solution directly to the
+    wells to stop the reaction.
+
+14. Measure the absorbance of each well at 450 nm on the spectrophotometer. Use the
+    spectrophotometer to generate a standard curve. If you don't see any color
+    change or dose response, your antibody concentration may have been too low. In
+    this case, use higher antibody concentrations. If the reaction is saturated and
+    you don't see any dose response, your antibody concentration may have been too
+    high. In this case, use lower antibody concentrations. The ideal range of the
+    standard curve will vary between targets and needs to be empirically determined.
+    The dilution series created in section 1, step 2 is a suggested starting point.
+    If your unknown sample's absorbance falls above the range of the standard curve
+    you will need to either include higher concentrations in the standard curve or
+    dilute the unknown sample (don't forget to factor in that dilution when
+    calculating the original sample concentration!).
+
+
+TIPS AND TROUBLESHOOTING
+================================================================================
+
+• The optimal concentration of primary antibody to use will vary and needs to be
+  empirically determined. If possible, run an initial test with serially diluted
+  primary antibody and determine the best concentration for your assay.
+
+• Washing steps can be done by hand but if available, use an automated ELISA plate
+  washer.
+
+
+SAMPLE DATA
+================================================================================
+
+The plate was coated with serial dilutions of human recombinant purified Desmin
+(blue) or a negative control protein, Apolipoprotein L3 (red) overnight,
+blocked, and incubated overnight with Anti-Desmin [D7]. Plates were incubated with
+HRP-linked anti-Human IgG secondary antibody for 2 hours, reacted with TMB
+substrate and the reaction stopped after 15 minutes. Absorbance was read at 450 nm
+and plotted against the micrograms of antigen loaded.
+
+Last reviewed on: October 8, 2024

biomni/tool/protocols/addgene/Addgene_ Colony Formation Titering Assay.txt

@@ -0,0 +1,221 @@
+PROTOCOL: Colony Formation Titering Assay
+SOURCE: https://www.addgene.org/protocols/colony-formation-titering-assay/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+This protocol can be used to titer lentivirus preparations that confer antibiotic
+resistance. Determining the titer of your lentiviral preparation allows you to
+control the multiplicity of infection (MOI) in downstream studies. This protocol
+was developed for A549 cells but can be adapted to a variety of target cell lines
+and selection markers. Note this assay requires staining resistant colonies with
+crystal violet solution and, therefore, the cells cannot be used for later
+experiments.
+
+
+WORKFLOW TIMELINE
+================================================================================
+
+Day 0: Seed and transduce cells
+
+Day 2: Replace media with fresh media containing selection reagent
+
+Days 3–14: Change media as needed
+
+Days 14–18: Stain cells and count resistant colonies
+
+
+EQUIPMENT
+================================================================================
+
+• Class II, Type A2 Biological Safety Cabinet
+• 0.5–10 µL single channel pipette
+• 2–20 µL single channel pipette
+• 20–200 µL single channel pipette
+• 200–1000 µL single channel pipette
+• Ice bucket
+• CO₂ incubator
+• Pipet controller
+• Hazardous waste container
+
+
+REAGENTS
+================================================================================
+
+• DMEM high glucose, Corning 10-013-CV
+• L-alanyl-L-glutamine (or alternative stable glutamine such as glutaGRO,
+  Corning 25-015-CI)
+• Heat-inactivated FBS
+• Polybrene (10 mg/mL), Millipore TR-1003-G
+• 1X PBS pH 7.4 without calcium or magnesium, Corning 21-040-CV (cations can
+  affect the attachment of adherent cells)
+• 0.1% Crystal Violet, Ward's Scientific CC0456-500ML
+• 0.22 µm luer-lock filter, Corning 431229
+• 10 mL luer-lock syringe, BD BD300912
+• 0.45 μm polyethersulfone filter, Nalgene, 565-0010 (for viral preps, if prep
+  was not previously filtered)
+• Microcentrifuge tubes, Neptune 3745.X
+• 6-well tissue culture treated dish, Corning 3516
+• 15 mL conical tubes, VWR 21008-216
+• Lentivirus preparation
+
+
+REAGENT PREPARATION
+================================================================================
+
+DMEM Complete: 10% v/v FBS and 4 mM L-alanyl-L-glutamine (or stable alternative,
+such as glutaGRO)
+
+To a 500 mL bottle of DMEM high glucose, add 55 mL of heat inactivated FBS and 5
+mL of 100X glutaGRO. Store at 4 °C.
+
+💡 PRO TIP: Different brands and lots of FBS can promote or inhibit
+transfection. Test a variety of brands and lots of FBS to find one suitable with
+your protocols. FBS can be purchased already heat-inactivated, or it can be
+inactivated in the lab by heating to 56 °C for 30 min.
+
+
+CONSIDERATIONS BEFORE YOU START
+================================================================================
+
+• The health of the target cell line is critical for obtaining accurate titers.
+
+• Check the cells for mycoplasma regularly.
+
+• Do not over or under-grow your cells.
+
+• Thaw a new vial of cells after 20–30 passages.
+
+• Do not add penicillin/streptomycin to the media, as it can affect viability and
+  transduction efficiency.
+
+• Titer will vary between cell lines.
+
+• It is not recommended that lentiviral supernatants be subjected to multiple
+  freeze-thaw cycles.
+
+• This protocol outlines the seeding of the cells at the same time as the
+  virus-mediated transduction. For some cell lines, transduction is optimized if
+  the cells are seeded the day before transduction. Note that in that case,
+  polybrene should only be added at the time of viral transduction, and not during
+  the cell seeding step.
+
+
+PROCEDURE
+================================================================================
+
+1. Before beginning a colony formation assay, the dose of antibiotic required to
+   kill your target cell line needs to be empirically determined. Treat the
+   target cells with a range of doses of antibiotic. Determine the minimum
+   concentration required to kill all of the cells. Use this dose for the colony
+   formation assay.
+
+2. Prepare a batch of DMEM complete containing 10 μg/mL polybrene by diluting 20
+   μL of 10 mg/mL polybrene into 20 mL of media.
+
+3. If using freshly collected virus, filter through a 0.45 μm polyethersulfone
+   filter to remove cells and debris.
+
+4. Lentiviral titer can decrease during cycles of freeze-thaw. If you are
+   freezing and aliquoting virus, it is recommended that you titer from the
+   frozen stock to account for any loss in titer associated with freeze-thaw.
+
+5. If using frozen virus, thaw the lentiviral aliquot on ice prior to use.
+
+6. Prepare 10-fold serial dilutions of the lentivirus into DMEM complete
+   containing 10 μg/mL polybrene:
+
+   Dilution      | Volume of Virus/Previous | Volume of DMEM Complete | Volume of Virus | Volume of Cells | Final Viral
+                 | Dilution (μL)            | + Polybrene (μL)       | Added (μL)     | Added (μL)     | Dilution
+   --------------|--------------------------|-------------------------|-----------------|----------------|------------
+   1:10          | 100 of Stock Virus       | 900                     | 150             | 1,350          | 1:100
+   1:100         | 100 of 1:10              | 900                     | 150             | 1,350          | 1:1,000
+   1:1,000       | 100 of 1:100             | 900                     | 150             | 1,350          | 1:10,000
+   1:10,000      | 100 of 1:1,000          | 900                     | 150             | 1,350          | 1:100,000
+   1:100,000     | 100 of 1:10,000         | 900                     | 150             | 1,350          | 1:1,000,000
+
+   Mix the dilutions well.
+
+   Note: the 1:10 dilution can usually be omitted because this dilution typically
+   produces a lawn of cells, which cannot be accurately quantified.
+
+7. Add 150 μL of a viral dilution to each well (each well gets one dilution).
+
+8. Also include an untransduced well by using 150 μL of media in place of virus.
+
+9. Seed 1,000 cells into each well of a 6-well dish.
+
+   Prepare a batch of cells as follows: Dilute 7,000 cells into 9.45 mL of DMEM
+   complete containing 10 μg/mL polybrene.
+
+10. Mix well by pipetting or inverting.
+
+11. Aliquot 1.35 mL of the cell suspension into each well of the 6-well dish
+    directly on top of the virus. Swirl gently to mix and place the dish in the
+    incubator.
+
+12. Incubate for 48–72 h.
+
+    💡 PRO TIP: Different cells may be more sensitive to lentivirus or to
+    polybrene, in which case this incubation step can be reduced to 8 h and the
+    media can be replaced with DMEM complete. However, antibiotic-containing media
+    should not be added until 48–72 h after transduction.
+
+13. Gently aspirate the media from the cells.
+
+14. Add 1.5 mL of DMEM complete containing the appropriate antibiotic.
+
+15. Incubate the cells for ~2 weeks. All of the cells in the untransduced
+    (negative) control well should be killed and no colonies should be visible.
+
+16. Every 3–4 days, gently aspirate the media and replace it with fresh DMEM
+    complete containing the appropriate antibiotic.
+
+17. When the untransduced (negative control) cells have died and colonies are
+    visible in the transduced wells, aspirate the media from the wells.
+
+18. Gently wash the cells with 1 mL of PBS and aspirate wash.
+
+19. Filter 0.1% crystal violet solution through a 0.22 μm filter to remove any
+    precipitates.
+
+20. Stain each well with 1 mL of 0.1% crystal violet for 10 min at RT.
+
+21. Gently remove the stain.
+
+22. Wash cells 3x with 1 mL of PBS, being careful not to disturb the colonies.
+
+23. Count the colonies for at least 2 of the dilutions.
+
+    💡 PRO TIP: The higher dilution wells tend to provide more accurate titers as
+    the likelihood of multiple integration events decreases as the dilution
+    increases.
+
+24. Calculate the transduction units per mL (TU/mL) as follows:
+
+    TU/mL = number of colonies / total volume in the well (mL) × dilution factor
+
+    e.g., If the 1:100,000 well has 75 colonies, then there are 75 colonies /
+    1.5 mL × 1,000,000 or 50,000,000 TU/mL
+
+    For a more accurate titer, take the average of multiple dilutions.
+
+
+SAMPLE DATA
+================================================================================
+
+A549 cells were transduced with the indicated serial dilutions of the lentiviral
+vector pRosetta. The following day, cells were treated with puromycin. Colonies
+were allowed to grow in the presence of selective reagent for 12 days with media
+exchanges every 3–4 days. Colonies were then stained with 0.1% crystal violet
+and counted.
+
+A549 cells were transduced with the indicated serial dilutions of the lentiviral
+vector pHAGE EF1α dCas9-KRAB. The following day, cells were treated with
+puromycin. Colonies were allowed to grow in the presence of selective reagent
+for 12 days with media exchanges every 3–4 days. After 12 days of selection, no
+cells have survived in the untransduced well, while lawns of cells are present in
+the lower dilutions, and single colonies are visible in the higher dilutions.
+
+Last reviewed on: August 31, 2023

biomni/tool/protocols/addgene/Addgene_ Coomassie Purity Stain of Recombinant Antibodies.txt

@@ -0,0 +1,307 @@
+PROTOCOL: Coomassie Purity Stain of Recombinant Antibodies
+SOURCE: https://www.addgene.org/protocols/coomassie-purity-stain-recombinant-antibodies/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+This protocol describes how to determine the purity and concentration of
+recombinant antibodies using ready-to-use bio-safe Coomassie G-250 stain
+(Addgene uses SimplyBlue SafeStain) and ImageJ software. The sample is
+separated by denaturing polyacrylamide gel electrophoresis alongside serial
+dilutions of a standard antibody of known concentration. After staining with
+Coomassie blue, protein band intensities are measured using ImageJ software and a
+standard curve is generated. The ratio of the antibody protein content to the
+total protein content of the sample is determined.
+
+Last Update: June 14, 2023
+
+
+WORKFLOW TIMELINE
+================================================================================
+
+Day 1: Run SDS-PAGE and stain gel
+
+Day 1 or later: Image analysis
+
+
+EQUIPMENT
+================================================================================
+
+• Heat block
+• 1–10 µL single channel pipette
+• 2–20 µL single channel pipette
+• 20–200 µL single channel pipette
+• 200–1000 µL single channel pipette
+• Pipette controller
+• Microcentrifuge
+• Electrophoresis chamber
+• Power supply
+• Rocking platform
+• Fume hood
+• Metal spatula
+• Razor blade
+• Plastic tray
+• Gel imaging system
+• ImageJ or similar software
+
+
+REAGENTS
+================================================================================
+
+• Purified recombinant antibody, 0.9–1.1 mg/mL
+• SimplyBlue SafeStain, Thermo Fisher, LC6060
+• 4-12% NuPage Novex bis-tris mini gel, 1mm thick, 10-well, Invitrogen NP0321BOX
+• 20X MOPS SDS running buffer, Invitrogen NP0001
+• 4X NuPage sample buffer, Invitrogen NP0007
+• 10X NuPage sample reducing agent (500 mM dithiothreitol, DTT), Invitrogen
+  NP0009
+• PageRuler Plus Prestained protein ladder, Invitrogen 26619
+• Pipette tips, 10 µL, VWR 76322-132
+• Pipette tips, 20 µL, VWR 76322-134
+• Pipette tips, 200 µL, VWR 76322-150
+• Pipette tips, 1000 µL, VWR 76322-154
+• Pipettes, 5 mL, VWR 89130-896
+• Pipettes, 10 mL, VWR 89130-898
+• Pipettes, 25 mL, VWR 89130-900
+• Pipettes, 50 mL, VWR 89130-902
+• Gel loading tips, Corning 4853
+• Microcentrifuge tubes, Neptune 3745.X
+• PBS, 1X pH 7.4, VWR 45000-446
+• 250 mL sterile bottles, Corning 430281
+• Deionized water
+• IgG isotype standard 2.5 mg/mL - can use commercial standard or validate an
+  in-house standard
+
+
+BEFORE STARTING
+================================================================================
+
+• Warm the hot plate to 100 °C.
+
+• Thaw IgG standard and prestained protein ladder on ice.
+
+
+REAGENT PREPARATION
+================================================================================
+
+Prepare IgG Standard
+
+Dilute the 2.5 mg/mL IgG isotype standard in PBS as follows:
+
+• 1.5 mg/mL: 18 µL of 2.5 mg/mL with 12 µL PBS
+• 1.0 mg/mL: 20 µL of 1.5 mg/mL with 10 µL PBS
+• 0.75 mg/mL: 15 µL of 1.0 mg/mL with 5 µL PBS
+• 0.5 mg/mL: 13.3 µL of 0.75 mg/mL with 6.7 µL PBS
+• 0.25 mg/mL: 10 µL of 0.5 mg/mL with 10 µL PBS
+• 0.125 mg/mL: 10 µL of 0.25 mg/mL with 10 µL PBS
+
+Transfer 5 µL of each standard to a microfuge tube.
+
+Add 8 µL of PBS to a total volume of 13 µL.
+
+Prepare the recombinant antibody
+
+1. Dilute or concentrate purified recombinant antibody to a concentration of
+   0.9–1.1 mg/mL.
+
+2. Transfer 5 µL of recombinant antibody to a microfuge tube.
+
+3. Add 8 µL of PBS to total 13 µL.
+
+Prepare 1X MOPS buffer
+
+Dilute 25 mL of 20X MOPS buffer into 475 mL of deionized water.
+
+Gently invert to mix.
+
+
+PROCEDURE
+================================================================================
+
+Section 1: SDS-PAGE
+
+1. Add 5 µL of 4X sample buffer to each sample.
+
+2. Add 2 µL 10X reducing agent to each sample.
+
+3. Spin the sample briefly in the microcentrifuge.
+
+4. Heat the samples for 10 min at 100 °C in a heat block.
+
+5. Spin the sample briefly in the microcentrifuge.
+
+6. Remove the gel from the plastic wrapper and rinse with deionized water.
+
+7. Gently remove the white sticker on the bottom of the gel.
+
+8. Place the gel in the electrophoresis chamber and secure it.
+
+   Note: Check the manufacturer's instructions if you are unsure of the correct
+   orientation of the gel.
+
+9. Carefully remove the comb from the gel.
+
+10. Rinse each well with 200 µL 1X MOPS running buffer.
+
+11. Fill the chamber with 1X MOPS running buffer.
+
+12. Make sure that the chamber is sealed.
+
+13. Load 5 µL of the prestained protein ladder to the appropriate well.
+
+    💡 PRO TIP: When possible, skip one lane before loading samples.
+
+14. Load 20 µL of each recombinant antibody sample to the appropriate well.
+
+    💡 PRO TIP: Leaving clear lanes between samples will make quantifying the gel
+    easier.
+
+15. Cover the electrophoresis chamber and attach to a power supply.
+
+16. Run the gel at 150 V for 1 h.
+
+    💡 PRO TIP: If the samples are running unevenly and the dye front looks like a
+    "smile", reduce the voltage.
+
+17. Turn off the power supply and unplug the electrophoresis chamber.
+
+18. Remove the gel from the chamber.
+
+19. Use the metal spatula to gently break the gel cast open.
+
+20. Use a razor blade to cut the top of the wells and bottom part of the gel where
+    dye is visible.
+
+Section 2: Staining the Gel
+
+1. Place the gel in a plastic tray with 100 mL of deionized water.
+
+2. Rinse gel with deionized water for 5 min with gentle agitation on a rocking
+   platform.
+
+3. Pour off the water in the sink.
+
+4. Add 20 mL of SimplyBlue SafeStain and incubate for 1 h with gentle agitation on
+   a rocking platform.
+
+5. Pour off the SimplyBlue SafeStain in the sink.
+
+6. Add 100 mL of deionized water and incubate for 1 h with gentle agitation on a
+   rocking platform.
+
+7. Pour off the water in the sink.
+
+8. Add 100 mL of deionized water and incubate for 1 h with gentle agitation on a
+   rocking platform.
+
+9. Pour off the water in the sink.
+
+10. Take a brightfield image of the gel with an appropriate imaging system.
+
+    Recombinant antibody preps should have 2 clear bands at ~50 kDa and ~25 kDa
+    corresponding to the heavy chain (HC) and light chain (LC) proteins,
+    respectively. There should be very little background staining.
+
+    💡 PRO TIP: A large shift in the electrophoretic mobility of the bands
+    (sample AR0016 in Figure 1) indicates that the samples may not have been
+    processed correctly in the cell (e.g., failure to cleave the signal peptide)
+    and may not be functional. This sample would fail Addgene QC and would not be
+    used.
+
+Section 3: Image Analysis
+
+Use ImageJ or a similar photo software to determine the relative intensity of the
+protein bands to the overall lane.
+
+1. Import the gel image into ImageJ.
+   • Select File.
+   • Select Open.
+   • Choose the location of the file to open.
+
+2. Change the image type to 8-bit.
+   • Select Image.
+   • Select Type.
+   • Choose 8-bit.
+
+3. Determine each lane of the gel as follows:
+   • Using the box tool, draw a box around the entire first gel lane.
+   • Select Analyze.
+   • Select Gels.
+   • Select Select First Lane.
+   • Drag the box to the next lane.
+   • Select Analyze.
+   • Select Gels.
+   • Select Select Next Lane.
+   • Repeat for all lanes.
+
+4. Plot the area under the curves for the protein bands as follows:
+   • Select Analyze.
+   • Select Gels.
+   • Select Plot Lanes.
+   • One graph per lane will appear with peaks representing each protein band.
+   • Use the line tool to connect the bottom of each peak.
+   • Select the wand tool.
+   • Fill in each peak by selecting the center of the peak.
+
+5. Export results as a csv file.
+   • In the results table, select File and Save As.
+
+6. Determine the percent purity of the samples:
+
+   P = ((H + L) / A) × 100
+
+   Where:
+   • P = Purity, %
+   • H = Area of HC peak
+   • L = Area of LC peak
+   • A = Total area of all bands in the lane
+
+   Example for AR0018 (lane 2 in Figure 1):
+   • Peak 1 (contaminant): 310.142
+   • Peak 2 (contaminant): 207.971
+   • Peak 3 (HC): 11469.296
+   • Peak 4 (LC): 4655.113
+   • Total Area: 16642.522
+   • HC + LC Area: 16124.409
+   • Purity: 96.887%
+
+   💡 PRO TIP: Addgene only uses recombinant antibody preps that have a purity
+   of ≥90% but some labs use unpurified tissue culture supernatant routinely
+   without any issues.
+
+7. Plot the mg/mL versus the HC + LC Area column for the IgG standards in Google
+   Sheets.
+   • Open exported CSV file.
+   • Select import.
+   • Choose file.
+   • Format as needed.
+   • Select the Insert tab, then Chart. This will add a blank chart to your
+     spreadsheet.
+   • In the Chart Editor, select the Chart Type drop down menu and select Scatter
+     chart.
+   • Change the data range and select the cells where the x-axis is represented.
+   • Add a linear trendline for the standards. Google Sheets can calculate this
+     for you as follows:
+     - In the Chart Editor, select the Customize tab.
+     - Select the Series drop down menu.
+     - Select the Trendline checkbox.
+
+8. Determine the linear equation and the R² of the trendline in Google Sheets as
+   follows:
+   • In the Customize tab, select the Label drop down menu and choose Use
+     Equation.
+   • Select the Show R² checkbox.
+
+   💡 PRO TIP: The R² of the trendline should be between 0.95–1.
+
+9. Use the equation of the trendline to determine the concentration of your
+   samples based on your sample's HC + LC area.
+
+   Example calculation:
+   If the sample HC + LC Area is 18179.442 and the linear equation of the
+   standard curve is y = 14725x + 3748.4, then substitute 18179.442 for y and
+   solve for x.
+   18179.442 = 14725x + 3748.4
+   x = 0.980
+   The concentration of the sample is 0.980 mg/mL

biomni/tool/protocols/addgene/Addgene_ Fluorescence Titering Assay.txt

@@ -0,0 +1,202 @@
+PROTOCOL: Fluorescence Titering Assay
+SOURCE: https://www.addgene.org/protocols/fluorescence-titering-assay/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+This protocol can be used to titer lentiviral preparations that carry a
+fluorescence marker. Determining the titer of your lentiviral vector allows you to
+control the multiplicity of infection (MOI) in downstream studies. This protocol
+was developed for 293T cells but can be adapted to a variety of target cell lines
+and selection markers. Note, because this protocol uses microscopy to determine
+the percentage of fluorescent cells, these cells can still be used in downstream
+applications.
+
+
+SAFETY WARNINGS
+================================================================================
+
+Lentivirus is generally considered biosafety level 2+. Please ensure that you are
+in compliance with your institution's biosafety regulations.
+
+
+WORKFLOW TIMELINE
+================================================================================
+
+Day 0: Seed 293T cells
+
+Day 1: Transduce cells
+
+Day 2 (am): Remove media, replace with fresh media
+
+Day 4–5: Count fluorescent cells
+
+
+EQUIPMENT
+================================================================================
+
+• Class II, Type A2 Biological Safety Cabinet
+• 0.5–10 µL single channel pipette
+• 2–20 µL single channel pipette
+• 20–200 µL single channel pipette
+• 200–1000 µL single channel pipette
+• Ice bucket
+• CO₂ incubator
+• Pipet controller
+• Hazardous waste container
+• Inverted fluorescence microscope
+
+
+REAGENTS
+================================================================================
+
+• DMEM high glucose, Corning 10-013-CV
+• L-alanyl-L-glutamine (or alternative stable glutamine such as glutaGRO,
+  Corning 25-015-CI)
+• Heat-inactivated FBS
+• Polybrene (10mg/mL), Millipore TR-1003-G
+• 1X PBS pH 7.4 without calcium or magnesium, Corning 21-040-CV (cations can
+  affect the attachment of adherent cells)
+• 0.45 μm polyethersulfone filter, Nalgene, 565-0010 (for viral preps, if prep
+  was not previously filtered)
+• Microcentrifuge tubes, Neptune 3745.X
+• 6-well tissue culture treated dish, Corning 3516
+• 15 mL conical tubes, VWR 21008-216
+• Lentivirus preparation
+
+
+REAGENT PREPARATION
+================================================================================
+
+DMEM Complete: 10% v/v FBS and 4 mM L-alanyl-L-glutamine (or stable alternative,
+such as glutaGRO)
+
+To a 500 mL bottle of DMEM high glucose, add 55 mL of heat-inactivated FBS and 5
+mL of 100X glutaGRO. Store at 4 °C.
+
+💡 PRO TIP: Different brands and lots of FBS can promote or inhibit
+transfection. Test a variety of brands and lots of FBS to find one suitable with
+your protocols. FBS can be purchased already heat-inactivated, or it can be
+inactivated in the lab by heating to 56 °C for 30 min.
+
+
+CONSIDERATIONS BEFORE YOU START
+================================================================================
+
+• The health of the target cell line is critical for obtaining accurate titers.
+
+• Check the cells for mycoplasma regularly.
+
+• Do not over- or under-grow your cells.
+
+• Thaw a new vial of cells after 20–30 passages.
+
+• Do not add penicillin/streptomycin to the media, as it can affect viability and
+  transduction efficiency.
+
+• Titer will vary between cell lines.
+
+• It is not recommended to subject lentiviral supernatants to multiple
+  freeze-thaw cycles.
+
+
+PROCEDURE
+================================================================================
+
+1. Seed 75,000 cells into each well of a 6-well dish.
+
+   Prepare a batch of cells as follows: Dilute 525,000 cells into 14 mL of DMEM
+   complete.
+
+2. Mix well by pipetting or inverting.
+
+3. Aliquot 2 mL of cell suspension into each well of the 6-well dish.
+
+4. Incubate the cells overnight.
+
+5. If using freshly collected virus, filter through a 0.45 μm polyethersulfone
+   filter to remove cells and debris.
+
+6. Lentiviral titer can decrease during cycles of freeze-thaw. If you are
+   freezing and aliquoting virus, it is recommended that you titer from the
+   frozen stock to account for any loss in titer associated with freeze-thaw.
+
+7. If using frozen virus, rapidly thaw the lentiviral aliquot on ice prior to
+   use.
+
+8. Prepare dilutions of the lentivirus into DMEM complete containing 10 μg/mL
+   polybrene. Note, this protocol was developed using low titer lentiviral vectors
+   (10⁵ TU/mL). If you anticipate that your viral stock will be higher titer,
+   consider additional dilutions.
+
+   Dilution    | Volume of Lentivirus | Volume of DMEM | Volume of 10mg/mL
+              | Stock (μL)           | complete (μL)  | polybrene (μL)
+   ------------|----------------------|----------------|------------------
+   1:10        | 150                  | 1348.5         | 1.5
+   1:25        | 60                   | 1438.5         | 1.5
+   1:50        | 30                   | 1468.5         | 1.5
+   1:75        | 20                   | 1478.5         | 1.5
+   1:100       | 15                   | 1483.5         | 1.5
+   Untransduced| 0                    | 1498.5         | 1.5
+
+9. Mix the dilutions well.
+
+10. Gently aspirate media from the cells.
+
+11. Add 1.5 mL of a viral dilution to each well (each well gets one dilution with
+    one well remaining as the untransduced control).
+
+12. Count the cells in the remaining well, a cell count is required for
+    calculating the titer.
+
+13. Incubate for 48–72 h.
+
+14. Gently aspirate media and replace with 1 mL of PBS.
+
+15. Calculate the fraction of fluorescent-positive cells in each well.
+
+    When calculating titer, only consider wells with less than 40% fluorescent
+    cells. Titering methods assume one integration event per cell. When the
+    percentage exceeds 40% you risk counting cells with multiple integration
+    events leading to underestimation of the true titer.
+
+16. Calculate the transduction units per mL (TU/mL) using either the dilution
+    factors (method 1) or the volume of virus (method 2):
+
+    Method 1: T = (N × F × D) / V_T
+
+    Where:
+    • T = Titer, TU/mL
+    • N = Number of cells transduced
+    • F = Fraction of cells with fluorescence
+    • D = Dilution Factor
+    • V_T = Transduction Volume, mL
+
+    For example: If 150,000 cells were transduced in the 1:100 well, resulting in
+    25% fluorescent cells, then the titer is:
+    (150,000 cells × 0.25 fluorescent × 100) / (1.5 mL) = 2.5×10⁶ TU/mL
+
+    Method 2: T = (N × F) / V_V
+
+    Where:
+    • T = Titer, TU/mL
+    • N = Number of cells transduced
+    • F = Fraction of cells with fluorescence
+    • V_V = Virus Volume, mL
+
+    For example: If 15 µL of virus added to 150,000 cells resulted in 25%
+    fluorescent cells, then the titer is:
+    (150,000 cells × 0.25 fluorescent) / (0.015 mL) = 2.5×10⁶ TU/mL
+
+    For a more accurate titer, take the average of multiple dilutions.
+
+
+SAMPLE DATA
+================================================================================
+
+293T cells were transduced with a range of dilutions of pHAGE-TO-dCas9-3XmCherry.
+72 h post transduction, cells were assayed for mCherry expression using
+fluorescence microscopy.
+
+Last reviewed on: September 9, 2023

biomni/tool/protocols/addgene/Addgene_ General Transfection.txt

@@ -0,0 +1,201 @@
+PROTOCOL: General Transfection
+SOURCE: https://www.addgene.org/protocols/transfection/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+This protocol describes a general method for transfecting mammalian cells using
+linear polyethylenimine. Transfections allow for transient expression of a gene
+of interest in a target cell line and can be useful for short term studies of
+protein function. We specifically use this protocol with a subclone of HEK293T
+cells optimized for viral production (AAVpro or Lenti-X), but any HEK293T cell
+line should work. This approach can be adapted for different cell lines and
+different transfection reagents.
+
+
+WORKFLOW TIMELINE
+================================================================================
+
+Day 0: Seed HEK293T cells (or a subclone of HEK293T optimized for viral
+       production)
+
+Day 1: Transfect Cells
+
+Day 2 (am): 18 h post transfection - Remove media, replace with fresh media
+
+Day 3 or more (am): Observe fluorescence, harvest cells, or perform your
+                    experiment
+
+
+EQUIPMENT
+================================================================================
+
+• Class II, Type A2 Biological Safety Cabinet
+• 0.5–10 µL single channel pipette
+• 2–20 µL single channel pipette
+• 20–200 µL single channel pipette
+• 200–1000 µL single channel pipette
+• Ice bucket
+• CO₂ incubator
+• Pipet controller
+• Hazardous waste container
+
+
+REAGENTS
+================================================================================
+
+• DMEM high glucose, Corning 10-013-CV
+• L-alanyl-L-glutamine (or alternative stable glutamine such as glutaGRO,
+  Corning 25-015-CI)
+• Heat-inactivated FBS
+• Low serum medium such as Opti-MEM or Opti-Pro SFM, Thermo Fisher, 12309019
+• 25 mM chloroquine
+• Polyethylenimine, linear MW 25,000 Da
+• Microcentrifuge tubes, Neptune 3745.X
+• 10 cm tissue culture dish, Corning 430167
+• Hydrochloric acid
+• Sodium hydroxide
+• 0.22 μm polyethersulfone (PES) filter
+• Syringes for filtering
+
+
+REAGENT PREPARATION
+================================================================================
+
+DMEM Complete: 10% v/v FBS and 4 mM L-alanyl-L-glutamine (or stable alternative
+such as glutaGRO)
+
+To a 500 mL bottle of DMEM high glucose, add 55 mL of heat-inactivated FBS and 5
+mL of 100X glutaGRO. Store at 4 °C.
+
+💡 PRO TIP: Different brands and lots of FBS can promote or inhibit
+transfection. Test a variety of brands and lots of FBS to find one suitable with
+your protocols. FBS can be purchased already heat inactivated or it can be
+inactivated in the lab by heating to 56 °C for 30 min.
+
+25 mM chloroquine diphosphate
+
+1. Dissolve 0.129 g of chloroquine diphosphate salt into 10 mL of sterile water.
+
+2. Filter sterilize through a 0.22 µm filter.
+
+3. Aliquot 50–100 µL and store at -20 °C.
+
+4. Aliquots can be thawed and stored at 4 °C prior to use. Thawed aliquots should
+   be discarded after 1–2 months.
+
+1 mg/mL polyethylenimine, linear MW 25,000 Da (PEI)
+
+1. Dissolve 100 mg of powder into 100 mL of deionized water.
+
+2. While stirring, slowly add hydrochloric acid until the solution clears.
+
+3. Check the pH of the solution. Use hydrochloric acid or sodium hydroxide to
+   adjust the pH to 7.0. Typically, the solution will be basic and will need
+   adjustment with hydrochloric acid first.
+
+   💡 PRO TIP: The pH of this solution will drift pretty rapidly upon addition of
+   acid or base. Add only a few drops at a time, allow them to mix and recheck the
+   pH to prevent over or undershooting the desired pH.
+
+4. Allow the solution to mix for 10 min and then recheck the pH to ensure that it
+   has not drifted.
+
+5. Filter the solution through a 0.22 µm membrane.
+
+6. Aliquot 500–1000 µL into sterile tubes.
+
+7. Store the tubes at -80 °C.
+
+8. After thawing the solution can be stored at 4 °C for up to 2 months. After 2
+   months, discard the tube and thaw a new working stock.
+
+
+CONSIDERATIONS BEFORE YOU START
+================================================================================
+
+• The health of the cell line is critical for obtaining high levels of virus.
+
+• HEK293T cells should be split 3 times a week:
+  - Monday: Plate 1x10⁶ cells in a 75 cm² flask in a volume of 15 mL.
+  - Wednesday: Plate 1x10⁶ cells in a T75 flask in a volume of 15 mL.
+  - Friday: Plate 8x10⁵ cells in a T75 flask in a volume of 15 mL.
+
+• Do not add antibiotics to the media.
+
+• Use cells that are below passage 20 for viral production.
+
+• The optimal mass DNA:mass PEI ratio will need to be empirically determined for
+  each new batch of 1 mg/mL PEI prepared.
+
+• There may be variation between batches of PEI depending on the user,
+  quantities of chemical used, volumes, pH adjustment etc. Consequently, each
+  batch needs to be validated and the best ratio of mass DNA:mass PEI determined.
+
+
+PROCEDURE
+================================================================================
+
+1. Seed HEK293T cells at 3.8x10⁶ cells per plate in DMEM complete in 10 cm
+   tissue culture plates.
+
+2. Incubate the cells at 37 °C, 5% CO₂ for ~20 h.
+
+3. Gently aspirate media, add 10 mL fresh DMEM complete containing 25 µM
+   chloroquine diphosphate and incubate ~5 h.
+
+   For 10 mL of DMEM complete, add 10 µL of 25 mM chloroquine diphosphate.
+
+4. Dilute 18.9 µg of DNA into 500 µL of Opti-Pro SFM.
+
+   💡 PRO TIP: Endotoxins can inhibit transfection; therefore, plasmid DNA
+   purification should include an endotoxin removal step. For high quality plasmid
+   DNA, the plasmid should also be propagated in an endonuclease negative E. coli
+   strain such as NEB stable.
+
+5. Dilute 1:3 (µg DNA:µg PEI) in 500 µL total of OptiPro SFM (per 10 cm plate).
+
+   56.7 µL of 1 mg/mL PEI, MW 25,000 Da in 443.3 µL of OptiPro SFM per 10 cm
+   plate
+
+   💡 PRO TIP: The ratio of µg DNA:µg PEI needs to be empirically determined.
+   Once a batch of PEI is prepared, transfect cells with a fluorescent plasmid
+   using a variety of ratios. Check the cells 1–2 days after transfection to
+   determine what ratio gives the highest percentage of GFP positive cells.
+
+   Refer to the table below for a possible range of ratios to test:
+
+   Ratio of DNA:PEI | Amount of DNA (μg) | Volume of 1 mg/mL PEI (μL)
+   -----------------|-------------------|---------------------------
+   1:1              | 18.9              | 18.9
+   1:2              | 18.9              | 37.8
+   1:3              | 18.9              | 56.7
+   1:4              | 18.9              | 75.6
+   1:5              | 18.9              | 94.5
+   1:6              | 18.9              | 113.4
+
+6. Gently add the diluted PEI to the diluted DNA. Add the diluted PEI dropwise
+   while gently flicking the diluted DNA tube. Incubate the mixture 15–20 min at
+   RT.
+
+7. Carefully transfer the transfection mix to your HEK293T cells. Add the
+   transfection mix dropwise, being careful not to dislodge the cells.
+
+8. Incubate the cells for 18 h, or until the following morning.
+
+9. The following morning, carefully aspirate the media. Replace the media with
+   15 mL of DMEM complete.
+
+10. Incubate the cells 24–48 h before checking for protein expression.
+
+
+SAMPLE DATA
+================================================================================
+
+Lenti-X 293T cells were transfected using 1:1, 1:2, 1:3 and 1:6 µg of pRosetta:µg
+of PEI. The 1:2 and 1:3 ratios provided high transfection efficiencies as can be
+seen here by the amount of green fluorescent protein expression (green in the
+right panels) with a limited effect on cell growth.
+
+Last reviewed on: November 7, 2023

biomni/tool/protocols/addgene/Addgene_ Kit Free RNA Extraction.txt

@@ -0,0 +1,266 @@
+PROTOCOL: Kit Free RNA Extraction
+SOURCE: https://www.addgene.org/protocols/kit-free-rna-extraction/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+This protocol describes how to extract RNA from tissues or cells without a kit. The
+general steps of this protocol include homogenization/lysis of cells or tissues,
+extraction of RNA, precipitation, and resuspension. This protocol provides two
+options: using Solution D (see reagent section for recipe) or using an all-in-one
+acid guanidinium thiocyanate-phenol solution such as TRIzol®. For the precipitation
+step, two options are also included: using Isopropanol or Lithium Chloride.
+
+This protocol was adapted from Chomczynski P and Sacchi N, 2006 and TRIzol® User
+Guide from ThermoFisher Scientific.
+
+
+BEFORE STARTING
+================================================================================
+
+RNA is not as stable as DNA and is susceptible to degradation by heat, RNases, and
+other enzymes.
+
+💡 PRO TIPS:
+
+• Always wear gloves, and whenever possible, keep RNA sample and reagents cold and
+  work quickly to reduce RNA degradation.
+
+• Keep work area, equipment, and reagents RNase-free (Use an RNase decontamination
+  solution, such as RNaseZap® or RNase AWAY®, may be used).
+
+• For more tips on working with RNA, read this blog post on RNA extraction without
+  a kit.
+
+Last Update: April 10, 2020
+
+
+EQUIPMENT
+================================================================================
+
+• Refrigerated microcentrifuge
+  (If you only have a non-refrigerated microcentrifuge, see step 4 of either
+  protocol before you start.)
+• Homogenizer
+• Vortexer
+• -20°C freezer
+• -80°C freezer
+
+
+REAGENTS
+================================================================================
+
+• Solution D (for Protocol Option #1):
+  - 4 M Guanidinium Thiocyanate
+  - 25 mM Sodium Citrate, pH 7.0
+  - 0.5% (wt/vol) N-laurosylsarcosine (Sarkosyl)
+  - 0.1 M 2-mercaptoethanol
+
+• TRIzol® or similar product such as TRI Reagent®, RNAzol®, QIAzol® (for Protocol
+  Option #2)
+
+• Water-saturated Phenol
+
+• 2 M Sodium Acetate pH 4
+
+• Chloroform/Isoamyl alcohol (49:1)
+
+• 75% Ethanol
+
+• RNase-free water or TE solution
+
+• RNase free tubes: microcentrifuge tubes, 4 mL polypropylene tubes
+
+• RNase decontamination solution like RNase AWAY® or RNaseZAP®
+
+• Isopropanol (for precipitation step, Option A)
+
+• 7.5 M Lithium Chloride (for precipitation step, Option B)
+
+• Glycogen (Optional)
+
+Caution: Make sure to read the SDS (Safety Data Sheet) for safety warnings and
+hazards for these reagents. Work in a well-ventilated space and under a fume hood
+when working with the volatile reagents in the list above.
+
+
+PROCEDURE
+================================================================================
+
+Option #1 - Solution D Protocol
+
+Before starting this protocol, make sure to prepare solution D (see reagent
+section above for the recipe). If using TRIzol®, jump down to the Option #2 -
+TRIzol® Protocol section below.
+
+1. Homogenize or lyse tissues or cells in Solution D.
+   • For tissues: use 1 mL of Solution D per 100 mg of cells.
+   • For cultured cells: use 1 mL of Solution D per 1 X 10⁷ cells.
+
+2. Allow sample(s) to sit at room temperature for 5 minutes to allow for
+   dissociation of the nucleoprotein complexes.
+
+   The effectiveness of your RNA isolation will depend on how effective your cell
+   lysis protocol is. While simple homogenization is effective for most mammalian
+   tissues; more hardy tissues such as bone, or bacteria/yeast/plant samples will
+   require additional steps to effectively lyse open the cells.
+
+3. Extract RNA from the homogenized sample(s). Transfer tissue/cell lysate to a 4
+   mL tube. Add the following sequentially to 1 mL of lysate:
+   • Add 0.1 mL of 2 M sodium acetate (pH 4.0), mix thoroughly by inversion.
+   • Add 1 mL water-saturated phenol, mix thoroughly by inversion.
+   • Add 0.2 mL of Chloroform/Isoamyl alcohol (49:1) and then shake vigorously by
+     hand for 10 seconds.
+
+4. Incubate sample(s) for 15 minutes on ice and centrifuge the sample(s) for 15
+   minutes at 12,000 × g at 4°C to separate RNA from the rest of the tissue/cell
+   lysate.
+
+   💡 PRO TIP: Having your samples spin at 4°C helps reduce RNA degradation. If
+   you don't have access to a refrigerated centrifuge, you can carefully bring a
+   centrifuge into a cold room for centrifugation. Once you're done using the
+   centrifuge, bring this equipment back to room temperature, as prolonged storage
+   in the cold room may damage it.
+
+5. Using a pipettor, transfer the top, aqueous phase to a new RNAse-free tube.
+
+   The mixture separates into a bottom organic layer, an interphase layer, and a
+   top, aqueous layer. Take care to not disturb or collect the interphase layer
+   with your pipette tip.
+
+   💡 PRO TIP: To collect as much of the top aqueous phase without disturbing the
+   interphase layer, consider using a lower volume pipettor like a p200 to collect
+   a majority of the aqueous phase. You may have to collect twice or more from the
+   same tube, but unlike using a p1000 tip it will give you more control of where
+   you're aiming your tip in the tube.
+
+6. Precipitate your sample(s). You can use either Isopropanol or Lithium Chloride
+   for this step.
+
+   Isopropanol (Option A): Add 1 volume of Isopropanol to the extracted aqueous
+   layer. Incubate at -20°C for 1 hour.
+
+   Lithium Chloride (Option B): LiCl selectively precipitates RNA versus DNA or
+   proteins. Add the correct amount of 7.5 M LiCl solution to bring the
+   concentration of LiCl in the extracted aqueous layer to 2.5 M. Incubate at
+   -20°C for 1 hour.
+
+   💡 PRO TIPS:
+   • If you anticipate your RNA yield to be small, RNase-free Glycogen may be used
+     as a carrier to facilitate RNA precipitation. This does not affect the quality
+     of RNA or downstream applications.
+   • To improve yield of RNA, instead of incubating at -20°C for 1 hour, you can
+     try incubating at -80°C overnight.
+
+7. Centrifuge for 20 minutes at 10,000 x g at 4°C and discard the supernatant.
+   There should be a gel-like white pellet of total RNA in the bottom of the tube.
+
+8. Wash the RNA by resuspending the pellet in 0.5–1 ml of 75% Ethanol and vortex
+   for a few seconds. Centrifuge for 5 minutes at 10,000 x g at 4 °C and remove
+   the supernatant.
+
+9. Remove as much of the ethanol wash as possible without disturbing the pellet.
+   Air-dry the pellet for 5-10 minutes.
+
+   Critical: It is important to not let the pellet get too dry before resuspending,
+   as this affects the solubility of the RNA.
+
+   💡 PRO TIP: To prevent overdrying, watch the pellet and carefully remove any
+   residual ethanol wash and add RNase-free water or TE as soon as the entire tube
+   is dried but while the white pellet is still visible.
+
+10. Resuspend RNA pellet in RNase-free water or TE. Quantify and assess the quality
+    of your RNA sample(s) using a spectrophotometer (such as a Nanodrop), agarose
+    gel, or bioanalyzer. For more information on nucleic acid quantification, see
+    our protocol for DNA quantification, which can be modified for RNA. Store your
+    RNA sample(s) at -80 °C to prevent RNA degradation and avoid multiple
+    freeze-thaw cycles.
+
+    💡 PRO TIP: To avoid multiple freeze-thaw cycles of your entire RNA sample,
+    consider making smaller aliquots of it and storing those in -80°C.
+
+Option #2 - TRIzol® Protocol
+
+1. Homogenize or lyse tissues or cells in TRIzol® or a similar product.
+   • For tissues: use 1 mL of TRIzol® per 100 mg of cells.
+   • For cultured cells: use 1 mL of TRIzol® per 1 X 10⁷ cells.
+
+2. Allow sample(s) to sit at room temperature for 5 minutes to allow for
+   dissociation of the nucleoprotein complexes.
+
+   The effectiveness of your RNA isolation will depend on how effective your cell
+   lysis protocol is. While simple homogenization is effective for most mammalian
+   tissues; more hardy tissues such as bone, or bacteria/yeast/plant samples will
+   require additional steps to effectively lyse open the cells.
+
+3. Extract RNA from the homogenized sample(s). Add 0.2 mL of Chloroform/Isoamyl
+   alcohol (49:1) per 1 mL of TRIzol® used. Shake vigorously by hand for 10
+   seconds.
+
+4. Incubate the sample(s) for 2-3 minutes on ice and centrifuge for 15 minutes at
+   12,000 × g at 4°C to separate RNA from the rest of the tissue/cell lysate.
+
+   💡 PRO TIP: Having your samples spin at 4°C helps reduce RNA degradation. If
+   you don't have access to a refrigerated centrifuge, you can carefully bring a
+   centrifuge into a cold room for centrifugation. Once you're done using the
+   centrifuge, bring this equipment back to room temperature, as prolonged storage
+   in the cold room may damage it.
+
+5. Using a pipettor, transfer the top, aqueous phase to a new RNAse-free tube.
+
+   The mixture separates into a bottom organic layer, an interphase layer, and a
+   top, aqueous layer. If using TRIzol®, the bottom layer will be a red-pink
+   color. Take care to not disturb or collect the interphase layer with your
+   pipette tip.
+
+   💡 PRO TIP: To collect as much of the top aqueous phase without disturbing the
+   interphase layer, consider using a lower volume pipettor like a p200 to collect
+   a majority of the aqueous phase. You may have to collect twice or more from the
+   same tube, but unlike using a p1000 tip it will give you more control of where
+   you're aiming your tip in the tube.
+
+6. Precipitate your sample(s). You can use either Isopropanol or lithium Chloride
+   for this step.
+
+   Isopropanol (Option A): Add 1 volume of Isopropanol to the extracted aqueous
+   layer. Incubate at -20°C for 1 hour.
+
+   Lithium Chloride (Option B): LiCl selectively precipitates RNA versus DNA or
+   proteins. Add the correct amount of 7.5 M LiCl solution to bring the
+   concentration of LiCl in the extracted aqueous layer to 2.5 M. Incubate at
+   -20°C for 1 hour.
+
+   💡 PRO TIPS:
+   • If you anticipate your RNA yield to be small, RNase-free Glycogen may be used
+     as a carrier to facilitate RNA precipitation. This does not affect the quality
+     of RNA or downstream applications.
+   • To improve yield of RNA, instead of incubating at -20°C for 1 hour, you can
+     try incubating at -80°C overnight.
+
+7. Centrifuge for 20 minutes at 10,000 x g at 4°C and discard the supernatant.
+   There should be a gel-like white pellet of total RNA in the bottom of the tube.
+
+8. Wash the RNA by resuspending the pellet in 0.5–1 ml of 75% Ethanol and vortex
+   for a few seconds. Centrifuge for 5 minutes at 10,000 x g at 4 °C and remove
+   the supernatant.
+
+9. Remove as much of the ethanol wash as possible without disturbing the pellet.
+   Air-dry the pellet for 5-10 minutes.
+
+   Critical: It is important to not let the pellet get too dry before resuspending,
+   as this affects the solubility of the RNA.
+
+   💡 PRO TIP: To prevent overdrying, watch the pellet and carefully remove any
+   residual ethanol wash and add RNase-free water or TE as soon as the entire tube
+   is dried but while the white pellet is still visible.
+
+10. Resuspend RNA pellet in RNase-free water or TE. Quantify and assess the quality
+    of your RNA sample(s) using a spectrophotometer (such as a Nanodrop), agarose
+    gel, or bioanalyzer. For more information on nucleic acid quantification, see
+    our protocol for DNA quantification, which can be modified for RNA. Store your
+    RNA sample(s) at -80 °C to prevent RNA degradation and avoid multiple
+    freeze-thaw cycles.
+
+    💡 PRO TIP: To avoid multiple freeze-thaw cycles of your entire RNA sample,
+    consider making smaller aliquots of it and storing those in -80°C.

biomni/tool/protocols/addgene/Addgene_ Lentivirus Production Protocol.txt

@@ -0,0 +1,224 @@
+PROTOCOL: Lentivirus Production Protocol
+SOURCE: https://www.addgene.org/protocols/lentivirus-production/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+This protocol can be used to produce lentivirus from a lentiviral vector
+transfected into 293T cells using a polyethylenimine (PEI) transfection protocol.
+This procedure can be modified for alternative packaging cell lines or
+transfection reagents. Once produced, lentivirus can be used for a variety of
+downstream applications such as stable-cell line generation.
+
+
+WORKFLOW TIMELINE
+================================================================================
+
+Day 0: Seed 293T packaging cells
+
+Day 1 (pm): Transfect packaging cells
+
+Day 2 (am): 18 h post-transfection. Remove media, replace with fresh media.
+
+Day 3-4 (am): Harvest virus
+
+
+EQUIPMENT
+================================================================================
+
+• Class II, Type A2 Biological Safety Cabinet
+• 0.5–10 µL single channel pipette
+• 2–20 µL single channel pipette
+• 20–200 µL single channel pipette
+• 200–1000 µL single channel pipette
+• Ice bucket
+• CO₂ incubator
+• Pipet controller
+• Hazardous waste container
+• pH meter
+• Stir plate
+• Magnetic Stir Bar
+
+
+REAGENTS
+================================================================================
+
+• DMEM high glucose, Corning 10-013-CV
+• L-alanyl-L-glutamine (or alternative stable glutamine such as glutaGRO,
+  Corning 25-015-CI)
+• Low serum medium such as Opti-MEM or Opti-Pro SFM, Thermo Fisher, 12309019
+• 25 mM chloroquine
+• Polyethylenimine, linear MW 25,000 Da
+• Heat-inactivated FBS
+• 1x PBS pH 7.4 without calcium or magnesium, Corning 21-040-CV (cations can
+  affect the attachment of adherent cells)
+• 0.45 μm polyethersulfone filter, Nalgene, 565-0010
+• Microcentrifuge tubes, Neptune 3745.X
+• 10 cm tissue culture dish, Corning 430167
+• 15 mL conical tubes, VWR 21008-216
+• Hydrochloric acid
+• Sodium hydroxide
+• 0.22 μm polyethersulfone (PES) filter
+• Transfection grade DNAs: Lentiviral Packaging plasmid(s), Envelope plasmid, and
+  Transfer plasmid encoding your insert of interest
+
+
+REAGENT PREPARATION
+================================================================================
+
+DMEM Complete (also known as D10): 10% v/v FBS and 4 mM L-alanyl-L-glutamine
+(or stable alternative, such as glutaGRO)
+
+To a 500 mL bottle of DMEM high glucose, add 55 mL of heat-inactivated FBS and 5
+mL of glutaGRO. Store at 4 °C.
+
+💡 PRO TIP: Different brands and lots of FBS can promote or inhibit
+transfection. Test a variety of brands and lots of FBS to find one suitable with
+your protocols. FBS can be purchased already heat inactivated or it can be
+inactivated in the lab by heating to 56°C for 30 min.
+
+1 mg/mL polyethylenimine, linear MW 25,000 Da (PEI)
+
+1. Dissolve 100 mg of powder in 100 mL of deionized water.
+
+2. While stirring, slowly add hydrochloric acid until the solution clears.
+
+3. Check the pH of the solution. Use hydrochloric acid or sodium hydroxide to
+   adjust the pH to 7.0. Typically, the solution will be basic and will need
+   adjustment with hydrochloric acid first.
+
+   💡 PRO TIP: The pH of this solution will drift rapidly upon addition of acid
+   or base. Add only a few drops at a time, allow them to mix and recheck the pH
+   to prevent over or undershooting the desired pH.
+
+4. Allow the solution to mix for 10 min and then recheck the pH to ensure that it
+   has not drifted.
+
+5. Filter the solution through a 0.22 μm membrane.
+
+6. Aliquot 500–1000 μL into sterile tubes.
+
+7. Store the tubes at -20 °C or -80 °C.
+
+8. After thawing, the solution can be stored at 4 °C for up to 2 months. After 2
+   months, discard the tube and thaw a new working stock.
+
+9. The optimal mass DNA:mass PEI ratio will need to be empirically determined for
+   each new batch of 1 mg/mL PEI and for each cell line.
+
+
+CONSIDERATIONS BEFORE YOU START
+================================================================================
+
+• The health of the packaging cell line is critical for obtaining high viral
+  titer.
+
+• 293T cells should be split 3 times a week:
+  - Monday: Plate 1×10⁶ cells in a T75 flask in 15 mL DMEM Complete.
+  - Wednesday: Plate 1×10⁶ cells in a T75 flask in 15 mL DMEM Complete.
+  - Friday: Plate 8×10⁵ cells in a T75 flask in 15 mL DMEM Complete.
+
+• Do not add pen-strep to the media.
+
+• Use cells that are below passage 15 for viral production.
+
+
+PROCEDURE
+================================================================================
+
+1. Seed 293T packaging cells at 3.8×10⁶ cells per plate in DMEM Complete in 10 cm
+   tissue culture plates.
+
+2. Incubate the cells at 37 °C, 5% CO₂ for ~20 h.
+
+3. Prepare a mixture of the three transfection plasmids into OptiPro SFM:
+
+   Reagent                    | Amount per 10 cm dish*
+   ---------------------------|------------------------
+   Packaging Plasmid (psPAX2) | 1.3 pmol
+   Envelope Plasmid (pMD2.G)  | 0.72 pmol
+   Transfer Plasmid*          | 1.64 pmol
+   OptiPro SFM to total       | 500 μL
+
+   *Plasmid concentrations and ratios should be optimized for each transfer
+   plasmid.
+
+   💡 PRO TIP: Endotoxins can inhibit transfection; therefore, plasmid DNA
+   purification should include an endotoxin removal step. For high quality plasmid
+   DNA, the plasmids should also be propagated in an endonuclease negative E.
+   coli strain such as NEB stable.
+
+4. Make a mixture of a total of 500 μL PEI-OptiPro SFM with enough PEI such that
+   the ratio of μg DNA:μg PEI is 1:3 (1000 μL total per 10 cm dish).
+
+   Using transfer plasmid pHAGE TRE dCas9-KRAB (total μg of plasmid DNA 27.8 μg),
+   this would be 83.4 μL of 1 mg/mL PEI in 416.6 μL of OptiPro SFM per 10 cm
+   dish.
+
+   💡 PRO TIP: There can be batch to batch variation when making the PEI working
+   stock, therefore the ratio of μg DNA:μg PEI needs to be empirically determined.
+   Once a batch of PEI is prepared, transfect cells with a fluorescent plasmid
+   using a variety of ratios. Check the cells 1-2 days after transfection to
+   determine what ratio gives the highest percentage of GFP positive cells.
+
+   Refer to the table below for a possible range of ratios to test:
+
+   Ratio of DNA:PEI | μg of DNA | μL of 1 mg/mL PEI
+   -----------------|-----------|------------------
+   1:1              | 18.9      | 18.9
+   1:2              | 18.9      | 37.8
+   1:3              | 18.9      | 56.7
+   1:4              | 18.9      | 75.6
+   1:5              | 18.9      | 94.5
+   1:6              | 18.9      | 113.4
+
+5. Gently add the diluted PEI mixture to the diluted DNA mixture. Add the diluted
+   PEI dropwise while gently flicking the diluted DNA tube. Incubate the mixture
+   12–15 min at RT.
+
+6. During the incubation, add 10 mL of DMEM Complete to a 15 mL conical and an
+   appropriate amount of 25 mM chloroquine to the conical so that the final
+   concentration in the dish (once transfection mixes are added) will be 25 uM of
+   chloroquine.
+
+7. After the incubation, add the DNA:PEI-Max mix to the conical containing the
+   media and chloroquine and mix well.
+
+8. Gently aspirate the media out of the previously seeded 10 cm plate.
+
+9. Slowly pipette the transfection mix onto the 10 cm plate, being careful not to
+   disturb the cells.
+
+10. Incubate the cells for 18 h, or until the following morning.
+
+11. The following morning, carefully aspirate the media. Replace the media with
+    10 mL of DMEM Complete or OptiPro SFM.
+
+12. Incubate the cells.
+
+13. Virus can be harvested at 48, 72, and 96 h post transfection in individual
+    harvests or a combined harvest where all the individual harvests are pooled.
+    If pooling harvests, transfer the harvested media to a polypropylene storage
+    tube and store at 4 °C between harvest.
+
+14. Centrifuge the viral supernatant at 2100 rcf for 5 minutes to pellet any
+    packaging cells that were collected during harvesting.
+
+15. Filter supernatant through a 0.45 μm PES filter.
+
+16. The viral supernatant can be stored at 4 °C for several hours but should be
+    aliquoted and snap frozen in liquid nitrogen and stored at -80 °C as soon as
+    possible to avoid loss of titer.
+
+
+SAMPLE DATA
+================================================================================
+
+293T cells were transfected with the GFP-expression plasmid pRosetta using μg
+total DNA to μg PEI ratios of 1:1, 1:2, 1:3 and 1:6. The 1:2 and 1:3 total
+DNA:PEI μg ratios provided high transfection efficiencies as measured by the
+highest proportion of GFP positive cells without limiting cell growth. Left
+panels: bright field images; right panels: GFP channel images.
+
+Last reviewed on: August 2, 2023

biomni/tool/protocols/addgene/Addgene_ Personal Protective Equipment (PPE) for BSL-1 and BSL-2 Labs.txt

@@ -0,0 +1,80 @@
+PROTOCOL: Personal Protective Equipment (PPE) for BSL-1 and BSL-2 Labs
+SOURCE: https://www.addgene.org/protocols/ppe/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+There are four biosafety levels. This protocol provides information for both
+biosafety level 1 (BSL-1) and biosafety level 2 (BSL-2). The BSL-1 classification is
+for labs working with low-risk microbes posing little to no threat of infection or
+disease in healthy humans. Many of the precautions are meant to protect individuals
+from potential accidents such as spills. BSL-2 is different because it includes
+labs that work with agents associated with diseases in healthy humans. BSL-2
+includes all of the precautions needed in BSL-1, however there are additional
+precautions that lie beyond PPE.
+
+
+EQUIPMENT
+================================================================================
+
+• Lab coat
+• Close-toed shoes
+• Gloves
+• Eye protection and face shields, as needed
+
+
+GUIDELINES
+================================================================================
+
+1. Your lab coat should be buttoned to protect your body and clothing from
+   potential spills.
+
+2. Gloves should fit well enough to be snug to your hand, but not so tight that
+   you cannot perform your duties. Be sure to try out different sizes to know your
+   fit. It is best to not reuse gloves. Dispose your gloves in a biohazard waste
+   container.
+
+3. Your lab coat and gloves should not be worn in "common areas" such as the lunch
+   room, offices, and bathroom. These items should remain in the laboratory. When
+   traveling between lab spaces, take your glove off of one hand and use your bare
+   hand to open doors.
+
+4. For those with latex allergies, be aware that some gloves may contain latex.
+   Consider nitrile gloves as an alternative.
+
+5. For extra precautions, tuck your sleeves into your gloves to prevent potential
+   spills down your sleeve. This may or may not be possible depending on the type
+   of sleeve.
+
+6. Lastly, don't touch your face!
+
+7. Close-toed shoes and long pants should be worn to protect your legs and feet
+   from potential spills. If wearing open-toed shoes, cloth coverings or
+   "booties" can be used to protect your feet.
+
+   💡 PRO TIP: If they are able, sometimes scientists work for a long time while
+   standing. Wear comfortable shoes that cushion your feet.
+
+8. Eye protection such as goggles and/or face shields can be used as needed. For
+   BSL-2 work always wear glasses/goggles in addition to the BSL-1 requirements.
+
+
+CONCLUSION
+================================================================================
+
+Although simple, following appropriate PPE protocol goes a long way. It is the
+easiest way to ensure that you're doing your work in the safest way possible.
+Protecting yourself is important!
+
+
+DISCLAIMER
+================================================================================
+
+The SOP presented here has been designed by the Addgene nonprofit plasmid
+repository and is being shared outside of Addgene for informational purposes only.
+If you choose to reuse or repurpose this SOP in another location, please note that
+you do so at your own risk; you should ensure that any local guidance is also
+adhered to. None of the authors, contributors, administrators, or anyone else
+associated with Addgene, can be held responsible for your use of the information
+contained in or linked to from these web pages.

biomni/tool/protocols/addgene/Addgene_ Pipetting Protocol.txt

@@ -0,0 +1,184 @@
+PROTOCOL: Pipetting Protocol
+SOURCE: https://www.addgene.org/protocols/pipetting/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+The pipette is an essential tool for nearly anyone in molecular biology. You'll use
+a pipette in many scenarios to accurately dispense small amounts of liquid (think:
+0.1 µL to 1 mL). When working in a laboratory, properly dispensing liquid ensures
+the accuracy of your experiment and any changes to the amount you are dispensing can
+negatively impact your experimental results. This protocol will guide you in
+selecting the correct pipette and pipette tip and using the pipette.
+
+Last Update: September 2022
+
+
+EQUIPMENT
+================================================================================
+
+• Pipettes
+• Pipette tips
+• Waste container
+• Containers to hold measured liquid (ex: microfuge tube, bottle, etc.)
+• Labels for containers
+
+
+REAGENTS
+================================================================================
+
+• Liquid for pipetting
+
+
+BACKGROUND INFORMATION
+================================================================================
+
+Anatomy of a Pipette
+
+This front view photo of a P1000 pipette shows the plunger at the top, which is
+used to draw and expel liquid into the pipette tip, the volume display which shows
+the set volume for the the pipette, and the tip ejector ring at the bottom which
+pushes the pipette tip off of the pipette.
+
+This side view photo of a P1000 pipette shows the volume adjustment ring right
+below the plunger. This ring changes the pipette volume. Below that is the tip
+ejector button which is used to push the tip ejector down the pipette to remove the
+tip.
+
+Choosing a Pipette and Pipette Tips
+
+Although there are many different types of pipettes that have different mechanisms
+to disperse liquids, this protocol is focused on single channel pipettes. These come
+in standard sizes: P2, P10, P20, P200, and P1000. The table shows the pipette with
+the appropriate dispense volume:
+
+Pipette    | Dispense Volume
+-----------|----------------
+P2         | 0.2 to 2 µL
+P10        | 1 to 10 µL
+P20        | 2 to 20 µL
+P100       | 10 to 100 µL
+P200       | 20 to 200 µL
+P1000     | 100 to 1,000 µL
+
+Each pipette should be used with the appropriate tips, which are made in a number of
+sizes and colors depending on the pipette they are used with and the volume to be
+dispensed. The boxes that the tips come in often indicate a volume range that the
+tip can hold. This should give you an idea of what pipette to use with it.
+
+The purpose of the pipette tip is so that the same pipette can be used for
+measuring different samples without cross contamination as long as the tip is
+changed between samples. Tips can come loose in a bag, or can come preloaded into
+tip boxes. In either case, once assembled into their boxes, pipette tips can be
+sterilized in an autoclave as needed to prevent contamination. When not in use, the
+tip box should be closed to prevent contamination.
+
+Reading the Volume on the Pipette
+
+Although each pipette performs the same function, the numbers are read differently
+on each type of pipette. This section will help you understand how to read each
+display to dispense the correct amount of liquid reagent.
+
+• P1000: The first red digit is thousands of µL, the middle is hundreds, and the
+  third is tens. Therefore, 1000 µL would read as 100 from top to bottom, while
+  650 µL would read as 065.
+
+• P200: The first digit is hundreds of µL, the second is tens, and the third is
+  ones. Therefore 100 µL would read as 100 from top to bottom, while 95 µL would
+  read as 095.
+
+• P20: The first digit is tens of µL, the second is ones, and the third (red)
+  digit is tenths. Therefore, 10 µL would read as 100, while 2.2 µL would read as
+  022.
+
+• P2: The first digit is ones of µL, the second red digit is tenths, and the
+  third (red) digit is hundredths. Therefore, 1 µL would read as 100, while 0.5 µL
+  would read as 050.
+
+Each volume display on these pipettes will also have small tick marks at the bottom.
+For the P1000, this represents the ones. For the P200, this represents the first
+decimal place. For the P20, this represents the second decimal place, and for the
+P2 this represents the third decimal place.
+
+
+PROCEDURE
+================================================================================
+
+How to Pipette
+
+Now that you know the different parts of the pipette, how to select tips, and how to
+read a pipette, you're ready to begin using the pipette. To the right is a photo of
+how you might hold a pipette.
+
+1. Without moving past the maximum possible setting on the pipette, set the desired
+   volume of the liquid on the pipette. Depending on the pipette you're using, you
+   will be rotating dials behind the numerical display or the top of the plunger of
+   the pipette.
+
+2. Select the pipette tip that is designed to fit your pipette. To load a tip onto
+   a pipette, open the tip box and push the end of the pipette onto the tip. Do not
+   touch the tips with your fingers to avoid contaminating the tip or puncturing
+   your glove. Ensure that the tip is properly set on the pipette to ensure
+   pipetting accuracy.
+
+3. Once the pipette is set and you have the liquid you wish to pipette ready, push
+   down the plunger. There will be multiple "stops" on the plunger, where you feel
+   resistance while pressing down on the pipette. Stop at the first "stop." The
+   first stop expels enough air from the pipette for an accurate measurement in the
+   next step. The second stop is meant to expel excess liquid from the tip once
+   you've reached step 8.
+
+4. Continuing to hold down the plunger, gently lower the pipette tip into the
+   liquid. Do not submerge the pipette itself into the liquid and avoid touching
+   the pipette to the inside of the tube. Only the tip should be in the liquid.
+
+   💡 PRO TIP: If pipetting from a relatively large container holding a small volume
+   of liquid, tilt the container holding the liquid so that it's easier for the
+   pipette tip to reach the liquid. Aim to let only the pipette tip into the
+   container in order to avoid contaminating the contents.
+
+5. Once the end of the tip is submerged, slowly release the plunger, allowing it to
+   go back to its resting position.
+
+6. Wait for the liquid to stop flowing into the pipette tip. Liquids of various
+   viscosity have different flow rates. The more viscous a liquid, the lower the
+   flow rate into the pipette tip.
+
+7. Once the liquid has finished drawing into the pipette tip, lift the pipette so
+   that the tip exits the liquid. When you lift the pipette tip out of the liquid,
+   no liquid should drip from the tip. This could indicate that the tip is not on
+   the pipette properly.
+
+8. Place the pipette tip into the container you are using to hold the measured
+   liquid (ex: another microcentrifuge tube, a bottle etc.). If there is already
+   other liquid in the receiving tube, place the pipette tip into the liquid. If
+   this is a new container, place the tip near the bottom of and gently in contact
+   with the container (capillary action between the container and the liquid will
+   help draw the liquid out of the pipette tip). Then, slowly press down on the
+   plunger all the way to the second stop to release the liquid.
+
+   💡 PRO TIP: Dispense until the first stop, wait, and then dispense to the second
+   stop to expel any residual liquid in the tip.
+
+9. Before releasing the plunger, lift the pipette tip out of the liquid. Once you
+   are clear of the liquid you can slowly release the plunger. Remove the entire
+   pipette from the container making sure not to touch the sides of the container
+   with the pipette.
+
+10. Discard the pipette tip by holding the pipette tip over your dedicated waste
+    container and pressing on the tip ejector button. If pipetting biohazardous
+    materials, be sure to discard into a biohazardous waste container.
+
+
+CONCLUSION
+================================================================================
+
+Pipetting accurately and precisely is important since even small changes in the
+volume could affect your experiments. Beyond knowing the proper way to pipette,
+it's important for scientists to calibrate their pipettes to make sure that they
+are properly tuned to dispense the correct amounts of liquid.
+
+If you are pipetting the same amount of liquid into different tubes or into wells
+of a microplate, you may consider using a multichannel pipette. Please watch
+Addgene's protocol on multichannel pipetting to learn more.

biomni/tool/protocols/addgene/Addgene_ Plasmid Cloning by Restriction Enzyme Digest (with Protocols).txt

@@ -0,0 +1,168 @@
+PROTOCOL: Plasmid Cloning by Restriction Enzyme Digest (with Protocols)
+SOURCE: https://www.addgene.org/protocols/subcloning/
+================================================================================
+
+SUMMARY
+================================================================================
+
+The following technique can be used to easily move any piece of DNA from one
+vector to another as long as it is already bounded by restriction sites that are
+also present in the same orientation on your target vector. If you are not sure
+what vector to use, you can check out our Empty Backbone Reference.
+
+
+BACKGROUND
+================================================================================
+
+Subcloning by restriction digest is a commonly used lab technique. For the
+purposes of this tutorial we will discuss how to move a cDNA from one plasmid to
+another. However, the same technique can be used to move promoters, selectable
+markers, or any other DNA element between plasmids.
+
+Let's assume that you are beginning a new project on your gene of interest (YGOI
+for short). You might need to express YGOI in cultured mammalian cells. The
+problem is that the only version of full-length cDNA you can find for YGOI is in a
+bacterial expression vector. Using subcloning, you can easily move YGOI into a
+mammalian expression vector.
+
+
+DESIGN (CHOOSING ENZYMES)
+================================================================================
+
+Many DNA analysis tools, including Addgene's Sequence Analyzer, allow you to
+identify which restriction sites are present in a given sequence. When selecting
+restriction enzymes, you want to choose enzymes that:
+
+• Flank your insert, but do not cut within your insert
+
+• Are in the desired location in your recipient plasmid (usually in the Multiple
+  Cloning Site (MCS)), but do not cut elsewhere on the plasmid
+
+• Will result in your insert being in the correct orientation in the recipient
+  plasmid. (You don't want to express the antisense version of your gene!)
+
+• Are in frame with tags or fusion proteins in the recipient plasmid (if you are
+  creating a fusion protein)
+
+Ideally, you will find two different restriction enzymes for your subcloning. It
+is also possible to use a single enzyme, but this will require phosphatase
+treatment of your recipient plasmid as well as a specifically designed test digest
+later to verify that the insert was cloned in the correct orientation.
+
+If you cannot find enzymes that meet these criteria, do not fear. You have other
+options, such as:
+
+Adding desired restriction sites to flank your insert: You can use PCR Based
+Cloning and add restriction sites to the ends of your oligos. This will allow
+you to produce a version of your insert flanked by restriction sites compatible
+with the recipient plasmid's MCS. However, you still need to avoid restriction
+enzymes that cut within your insert.
+
+Adding desired restriction sites to your recipient plasmid: You can modify the
+MCS of your recipient plasmid using Annealed-oligo Cloning.
+
+If you are lucky enough to have multiple options for enzymes that flank your
+insert and will result in correct orientation in the recipient plasmid, it is
+useful to see if one set of enzymes will work in the same restriction enzyme
+buffer (see New England Biolabs for more information about restriction enzyme
+buffers). If you select enzymes that can function in the same buffer, it will
+save you time in future steps.
+
+
+EXPERIMENTAL PROCEDURE
+================================================================================
+
+1. Digest your DNA
+
+   Set up restriction digests for your donor and recipient plasmids. Because you
+   lose some DNA during the gel purification step, it is important to digest plenty
+   of starting material. We recommend 1.5-2μg of donor plasmid and 1μg of
+   recipient plasmid. It is also critical that as much of the recipient plasmid as
+   possible be cut with both enzymes, and therefore it is important that the
+   digest go at least 4 hours and as long as overnight.
+
+   If you are going to use only one restriction enzyme, or enzymes that have
+   compatible overhangs or no overhangs after digestion, you will need to use a
+   phosphatase to prevent re-circularization of the recipient plasmid. You should
+   treat your digested recipient plasmid with a phosphatase prior to the ligation
+   step or prior to the gel purification step, depending on the phosphatase you
+   choose. CIP (calf alkaline phosphatase) or SAP (shrimp alkaline phosphatase) are
+   commonly used. Follow the manufacturer's instructions.
+
+2. Isolate your insert and vector by gel purification
+
+   Run your digested DNA on an agarose gel and conduct a gel purification to
+   isolate the DNA. When running a gel for purification purposes it is important
+   to have nice crisp bands and to have space to cut out the bands. Because of
+   this we recommend that you use a wide gel comb, run the gel on the slower side,
+   and skip lanes between samples. In addition to a DNA ladder standard, it is
+   also a good idea to run an uncut sample of each plasmid to help with
+   troubleshooting if your digests don't look as you expected.
+
+   Once you have cut out and purified your insert and recipient plasmid backbone
+   bands away from the gel via your favorite gel purification method, it is
+   important to determine the concentration of recovered DNA.
+
+3. Ligate your insert into your vector
+
+   Conduct a DNA Ligation to fuse your insert to your recipient plasmid.
+
+   We recommend around 100ng of total DNA in a standard ligation reaction. You
+   ideally want a recipient plasmid to insert ratio of approximately 1:3. Since
+   the number of base pairs for each varies, it is difficult to calculate this
+   based on DNA concentration alone. One method is to conduct 2 ligations for each
+   plasmid you are trying to create, with varying ratios of recipient plasmid to
+   insert.
+
+   It is also important to set up negative controls in parallel. For instance, a
+   ligation of the recipient plasmid DNA without any insert will tell you how much
+   background you have of uncut or self-ligating recipient plasmid backbone.
+
+4. Transformation
+
+   Transform your ligation reaction into your bacterial strain of choice. Follow
+   the manufacturer's instructions for your competent cells.
+
+   For most standard cloning, you can transform 1-2μl of your ligation reaction
+   into competent cells such as DH5alpha or TOP10. If using much less total DNA
+   (<1ng) or if you are having trouble getting colonies, you might want to use
+   higher competency cells. Additionally, if your final product is going to be very
+   large (>10kb) you might want to use electro-competent cells instead of the
+   more common chemically-competent cells.
+
+   The number of bacterial colonies resulting from your transformation will give
+   you the first indication as to whether your transformation worked. Your recipient
+   plasmid + insert plate should have significantly more colonies than the
+   recipient plasmid alone plate. The recipient plasmid alone control will tell
+   you your "background" level or more specifically it will tell you how many
+   colonies you can expect on your recipient plasmid + insert plate that are not
+   correct.
+
+   If you have a high number of colonies on your recipient plasmid alone plate, you
+   can try ligating the recipient plasmid alone in the presence and absence of
+   ligase. If the colonies are a result of uncut empty plasmid, you will still have
+   colonies when you do not add ligase. If the colonies are a result of recipient
+   plasmid self-ligation, you will see significantly more colonies when you add
+   ligase.
+
+   If you do not see any colonies, you should conduct a positive control to ensure
+   that your transformation worked. You should also verify that you are plating on
+   the appropriate antibiotic and try varying the amount of recipient plasmid to
+   insert in the ligation reaction.
+
+5. Isolate the Finished Plasmid
+
+   Finally, you will need to pick individual bacterial colonies and check them for
+   successful ligations. Pick 3-10 colonies depending on the number of background
+   colonies on your control plate (the more background, the more colonies you will
+   need to pick) and grow overnight cultures for DNA purification.
+
+   After purifying the DNA, conduct a diagnostic restriction digest of 100-300ng
+   of your purified DNA with the enzymes you used for the cloning. Run your digest
+   on an agarose gel. You should see two bands, one the size of your vector and
+   one the size of your new insert. If you used only one enzyme or used enzymes
+   with compatible overhangs you will need to verify the orientation of your
+   insert, so you may want to design a diagnostic digest for this purpose.
+
+Congratulations, you now have YGOI in a mammalian expression vector and can
+begin your studies.

biomni/tool/protocols/addgene/Addgene_ Protocol - Bacterial Transformation.txt

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+PROTOCOL: Bacterial Transformation
+SOURCE: https://www.addgene.org/protocols/bacterial-transformation/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+Transformation is the process by which foreign DNA is introduced into a cell.
+Transformation of bacteria with plasmids is important not only for studies in
+bacteria but also because bacteria are used as the means for both storing and
+replicating plasmids. Because of this, nearly all plasmids (even those designed
+for mammalian cell expression) carry both a bacterial origin of replication and
+an antibiotic resistance gene for use as a selectable marker in bacteria.
+
+Scientists have made many genetic modifications to create bacterial strains that
+can be more easily transformed and that will help to maintain the plasmid without
+rearrangement of the plasmid DNA. Additionally, specific treatments have been
+discovered that increase the transformation efficiency and make bacteria more
+susceptible to either chemical or electrical based transformation, generating what
+are commonly referred to as 'competent cells.'
+
+Many companies sell competent cells, which come frozen and are prepared for optimal
+transformation efficiencies upon thawing.
+
+For the highest transformation efficiency, we recommend that you follow the
+instructions that came with your competent cells.
+
+💡 PRO TIPS:
+
+• Commercial competent cells range significantly in their transformation
+  efficiency. The lowest efficiency cells (usually the least expensive) are fine
+  for transforming plasmid DNA for the purposes of storage and amplification.
+  Higher efficiency cells are more important if you will be transforming with very
+  small amounts of DNA or if you're multiple plasmids at once.
+
+• To save money, many labs also make their own competent cells. This is a
+  relatively simple procedure and is useful for performing low efficiency
+  transformations.
+
+Last Update: Nov. 13, 2017
+
+
+EQUIPMENT
+================================================================================
+
+• Shaking incubator at 37 °C
+• Stationary incubator at 37 °C
+• Water bath at 42 °C
+• Ice bucket filled with ice
+• Microcentrifuge tubes
+• Sterile spreading device
+
+
+REAGENTS
+================================================================================
+
+• LB agar plate (with appropriate antibiotic)
+• LB or SOC media
+• Competent cells
+• DNA you'd like to transform
+
+
+PROCEDURE
+================================================================================
+
+1. Take competent cells out of -80°C and thaw on ice (approximately 20-30 mins).
+
+2. Remove agar plates (containing the appropriate antibiotic) from storage at 4°C
+   and let warm up to room temperature and then (optional) incubate in 37°C
+   incubator.
+
+3. Mix 1 - 5 μl of DNA (usually 10 pg - 100 ng) into 20-50 μL of competent cells
+   in a microcentrifuge or falcon tube. GENTLY mix by flicking the bottom of the
+   tube with your finger a few times.
+
+   💡 PRO TIP: Transformation efficiencies will be approximately 10-fold lower
+   for ligation of inserts to vectors than for an intact control plasmid.
+
+4. Incubate the competent cell/DNA mixture on ice for 20-30 mins.
+
+5. Heat shock each transformation tube by placing the bottom 1/2 to 2/3 of the
+   tube into a 42°C water bath for 30-60 secs (45 secs is usually ideal, but this
+   varies depending on the competent cells you are using).
+
+6. Put the tubes back on ice for 2 min.
+
+7. Add 250-1,000 μl LB or SOC media (without antibiotic) to the bacteria and grow
+   in 37°C shaking incubator for 45 min.
+
+   💡 PRO TIP: This outgrowth step allows the bacteria time to generate the
+   antibiotic resistance proteins encoded in the plasmid backbone so that they will
+   be able to grow once plated on the antibiotic containing agar plate. This step
+   is not critical for Ampicillin resistance but is much more important for other
+   antibiotic resistances.
+
+8. Plate some or all of the transformation onto a 10 cm LB agar plate containing
+   the appropriate antibiotic.
+
+   💡 PRO TIPS:
+   • We recommend that you plate 50 μL on one plate and the rest on a second plate.
+     This gives the best chance of getting single colonies, while allowing you to
+     recover all transformants.
+   • If the culture volume is too big, gently collect the cells by centrifugation
+     and resuspend in a smaller volume of LB so that there isn't too much liquid
+     media on the agar plates. If the agar plate doesn't dry adequately before the
+     cells begin dividing, the bacteria diffuse through the liquid and won't grow
+     in colonies.
+
+9. Incubate plates at 37°C overnight.
+
+
+TIPS AND FAQ
+================================================================================
+
+How can I save time when carrying out transformations?
+
+If you are not concerned with transformation efficiency (such as when you have a
+tube of plasmid DNA and just need to transform bacteria so that you can grow up
+more of the plasmid) you can save a lot of time by shortening or skipping many
+steps and will still get enough colonies for your next step. Remember that each of
+these shortcuts will reduce the efficiency of the transformation, so when higher
+efficiency is needed follow the complete protocol.
+
+• Thaw the competent cells in your hand instead of on ice
+• Reduce step 4 from 20 - 30 mins to 2 mins on ice before heat-shock
+• Shorten or skip the outgrowth (for Ampicillin resistance it is ok to completely
+  skip the outgrowth, for the other antibiotics it is a good idea to outgrow for at
+  least 20-30 mins)
+
+I get very few if any transformants when transforming large plasmids (>10 kb) or
+BACs, what can I do?
+
+Chemically competent cells are fast and easy to use, but are less efficient at
+taking up larger plasmids. If you need to transform large plasmids, it is a good
+idea to use electro-competent cells. Instead of relying on the heat-shock to cause
+the cells to take up the DNA, an electro-magnetic field is applied to the
+cell/DNA mixture to induce membrane permeability. To do this you will need to
+have access to an electroporator and the appropriate cuvettes. Follow the
+manufacturer's instructions for each.
+
+I got no transformants. What went wrong?
+
+Check that you are plating on an LB Agar plate containing the correct antibiotic.
+The resistance gene on your plasmid must match the antibiotic on the plate. You
+should also add a positive control (many companies include a positive control
+plasmid with their competent cells) to ensure that your transformation procedure
+is working.
+
+TIP: Sometimes less is more.
+
+Although it may be counter-intuitive, you will often get higher transformation
+efficiencies with less DNA, especially when using highly competent cells. If you
+used 100-1000 ng of total DNA in a ligation you will often get more colonies if
+you use 1 μl of a 1:5 or 1:10 dilution rather than 1 μl directly.

biomni/tool/protocols/addgene/Addgene_ Protocol - How to Design Primers.txt

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+PROTOCOL: How to Design Primers
+SOURCE: https://www.addgene.org/protocols/primer-design/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+Oligonucleotide primers are necessary when running a PCR reaction. One needs to
+design primers that are complementary to the template region of DNA. They are
+synthesized chemically by joining nucleotides together. One must selectively block
+and unblock repeatedly the reactive groups on a nucleotide when adding a nucleotide
+one at a time. The main property of primers is that they must correspond to
+sequences on the template molecule (must be complementary to template strand).
+However, primers do not need to correspond to the template strand completely; it is
+essential, however, that the 3' end of the primer corresponds completely to the
+template DNA strand so elongation can proceed. Usually a guanine or cytosine is
+used at the 3' end, and the 5' end of the primer usually has stretches of several
+nucleotides. Also, both of the 3' ends of the hybridized primers must point toward
+one another.
+
+The size of the primer is very important as well. Short primers are mainly used
+for amplifying a small, simple fragment of DNA. On the other hand, a long primer
+is used to amplify a eukaryotic genomic DNA sample. However, a primer should not be
+too long (> 30-mer primers) or too short. Short primers produce inaccurate,
+nonspecific DNA amplification product, and long primers result in a slower
+hybridizing rate. On average, the DNA fragment that needs to be amplified should be
+within 1-10 kB in size.
+
+The structure of the primer should be relatively simple and contain no internal
+secondary structure to avoid internal folding. One also needs to avoid
+primer-primer annealing which creates primer dimers and disrupts the amplification
+process. When designing, if unsure about what nucleotide to put at a certain
+position within the primer, one can include more than one nucleotide at that
+position termed a mixed site. One can also use a nucleotide-based molecular insert
+(inosine) instead of a regular nucleotide for broader pairing capabilities.
+
+
+PRIMER PROPERTIES
+================================================================================
+
+Taking into consideration the information above, primers should generally have the
+following properties:
+
+• Length of 18-24 bases
+• 40-60% G/C content
+• Start and end with 1-2 G/C pairs
+• Melting temperature (Tm) of 50-60°C
+• Primer pairs should have a Tm within 5°C of each other
+• Primer pairs should not have complementary regions
+
+Note: If you will be including a restriction site at the 5' end of your primer,
+note that a 3-6 base pair "clamp" should be added upstream in order for the enzyme
+to cleave efficiently (e.g. GCGGCG-restriction site-your sequence).

biomni/tool/protocols/addgene/Addgene_ Protocol - How to Inoculate a Bacterial Culture.txt

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+PROTOCOL: How to Inoculate a Bacterial Culture
+SOURCE: https://www.addgene.org/protocols/inoculate-bacterial-culture/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+Plasmids can carry one or more antibiotic resistance genes, which confer
+resistance to a specific antibiotic to the bacteria carrying them. The presence of
+an antibiotic resistance gene on a plasmid allows researchers to easily isolate
+bacteria containing that plasmid from bacteria that do not contain it by
+artificial selection (i.e., growing the bacteria in the presence of the
+antibiotic).
+
+Luria broth (LB) is a nutrient-rich media commonly used to culture bacteria in
+the lab. LB agar plates are frequently used to isolate individual (clonal)
+colonies of bacteria carrying a specific plasmid. However, a liquid culture is
+capable of supporting a higher density of bacteria and is used to grow up
+sufficient numbers of bacteria necessary to isolate enough plasmid DNA for
+experimental use. The following protocol is for inoculating an overnight culture
+of liquid LB with bacteria.
+
+
+PROCEDURE
+================================================================================
+
+1. Prepare liquid LB. For example, to make 400 mL of LB, weigh out the following
+   into a 500 mL glass bottle:
+   • 4 g NaCl
+   • 4 g Tryptone
+   • 2 g Yeast Extract
+   • and dH₂O to 400 mL
+
+   Note: If your lab has pre-mixed LB agar powder, use the suggested amount,
+   instead of the other dry ingredients above.
+
+2. Loosely close the cap on the bottle (do NOT close all the way or the bottle may
+   explode!) and then loosely cover the entire top of the bottle with aluminum
+   foil. Autoclave and allow to cool to room temperature. Now screw on the top of
+   the bottle and store the LB at room temperature.
+
+3. When ready to grow your culture, add liquid LB to a tube or flask and add the
+   appropriate antibiotic to the correct concentration (see Preparing Antibiotics
+   below).
+
+   Note: If you intend to do a miniprep, you will usually want to start 2 mL in a
+   Falcon tube. For larger preps, you might want to use as much as a liter of LB
+   in a 2 L Erlenmeyer flask.
+
+4. Using a sterile pipette tip or toothpick, select a single colony from your LB
+   agar plate.
+
+5. Drop the tip or toothpick into the liquid LB + antibiotic and swirl.
+
+   Note: You can also use a long toothpick to select the colony, swirl it in the
+   LB, and remove prior to incubating.
+
+6. Loosely cover the culture with sterile aluminum foil or a cap that is not air
+   tight. Alternatively, you could use vented bioreactor tubes, dual position
+   snap-cap tubes, or permeable film to allow for aeration.
+
+7. Incubate bacterial culture at 37 °C for 12–18 h in a shaking incubator.
+
+   Note: Some plasmids or strains require growth at 30 °C. If so, you will likely
+   need to grow for a longer time to get the correct density of bacteria since
+   they will grow more slowly at lower temperatures.
+
+8. After incubation, check for growth, which is characterized by a cloudy haze in
+   the media (Figure 1).
+
+   Notes:
+   • Some protocols require bacteria to be in the log phase of growth. Check the
+     instructions for your specific protocol and conduct an OD600 to measure the
+     density of your culture if needed.
+   • A good negative control is LB media + antibiotic without any bacteria
+     inoculated. You should see no growth in this culture after overnight
+     incubation.
+
+9. Optional: For long term storage of the bacteria, you can proceed with creating
+   a glycerol stock.
+
+10. You can now isolate your plasmid DNA from the bacterial culture by following
+    the Isolating Your Plasmid DNA protocol.
+
+
+PREPARING ANTIBIOTICS
+================================================================================
+
+Create a stock solution of your antibiotic. Unless otherwise indicated, the
+antibiotic powder can be dissolved in dH₂O. Addgene recommends making 1000X stock
+solutions and storing aliquots at -20 °C.
+
+To use, dilute your antibiotic into your LB medium at 1:1,000. For example, to
+make 100 mL of LB/ampicillin growth media, add 100 μL of a 100 mg/mL ampicillin
+stock (1000X stock) to 100 mL of LB.
+
+Antibiotic Concentrations:
+
+Commonly Used Antibiotics | Recommended Concentration
+--------------------------|-------------------------
+Ampicillin                | 100 µg/mL
+Bleocin                   | 5 µg/mL
+Carbenicillin             | 100 µg/mL
+Chloramphenicol           | 25 µg/mL
+Coumermycin               | 25 µg/mL
+Gentamycin                | 10 µg/mL
+Kanamycin                 | 50 µg/mL
+Spectinomycin             | 50 µg/mL
+Tetracycline              | 10 µg/mL
+
+
+TIPS AND FAQ
+================================================================================
+
+What is the difference between high copy and low copy plasmids?
+
+The copy number refers to the number of copies of an individual plasmid within a
+single bacterial cell (Figure 2). Large plasmids usually have a low copy number
+(approximately one or two copies per cell) and they need to grow for longer
+periods of time (approximately 18–30 h). On the other hand, smaller plasmids can
+be present in large numbers, 50 or more per cell, and have a high copy number.
+High copy number plasmids should only need to be grown for 12–16 h on average.
+Certain features of a plasmid may render it low copy regardless of plasmid size.
+See the plasmid's info page to determine if your plasmid is high or low copy.
+
+I didn't get any growth after overnight incubation. What went wrong?
+
+• Try growing the culture for more time. Some bacterial cultures grow more slowly.
+  Also, bacteria incubated at 30 °C rather than 37 °C often require longer
+  incubation times.
+
+• Double check that the antibiotic in your LB media matches the antibiotic
+  resistance on your plasmid.
+
+• If the bacteria on your LB agar plates are not fresh, you should streak your
+  bacteria onto a new LB agar plate before growing in liquid culture.
+
+• More aeration may help to increase the density of the culture. Normally cultures
+  shake at 150–250 rpm, increase this to 350–400 rpm to obtain a higher cell
+  density.

biomni/tool/protocols/addgene/Addgene_ Protocol - How to Perform Sequence Analysis.txt

@@ -0,0 +1,108 @@
+PROTOCOL: How to Perform Sequence Analysis
+SOURCE: https://www.addgene.org/protocols/sequence-analysis/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+Sequence verification of important plasmid features (such as the gene/insert,
+fusion proteins, point mutations, deletions, etc.) involves selecting one or more
+unique oligonucleotide primers that flank the regions of your plasmid that you
+wish to confirm. Addgene's plasmid information pages recommend 5' and 3'
+sequencing primers. These primers typically anneal to the backbone and can help
+you verify the ends of the insert. Longer genes may require internal primers that
+anneal to a specific region of the insert.
+
+You can find Addgene's sequencing results by clicking on the "View Sequences" link
+on the Plasmid Information Pages. Addgene sequences the plasmid to verify tags,
+mutations and a portion of the insert, but we do not sequence the entire plasmid.
+Addgene strongly recommends that you sequence any portion of the plasmid that is
+important for your experiments.
+
+
+FACTORS TO CONSIDER WHEN SELECTING SEQUENCING PRIMERS
+================================================================================
+
+Is the primer unique?
+
+Make sure the primer only anneals once within the entire construct.
+
+Is the primer an appropriate distance from the target?
+
+The primer should be a minimum of 50 nucleotides and a maximum of 300 nucleotides
+from your target.
+
+Are there commercially available primers that can be used for sequencing?
+
+There are many widely used, common primers that are often found within the
+plasmid backbone (See Addgene's Sequencing Primers for reference). Many
+sequencing cores have a list of common primers that can be requested for no
+additional charge.
+
+What primers were used by Addgene during quality control?
+
+Addgene lists the primers used to obtain each result above the posted sequence in
+the "View Sequence" link.
+
+Is your custom primer found within the Addgene sequencing results?
+
+If you require additional sequencing and need to design a custom primer, Addgene
+recommends using Addgene's sequencing results as a reference for primer design.
+
+
+SEQUENCING RESULTS
+================================================================================
+
+A good sequencing reaction will produce between 300-900 base pairs of useable
+sequence. You should receive your sequencing results as a trace file (.ab1) which
+graphically depicts the sequence as a series of colored peaks corresponding to
+one of the four nucleotide bases. This is an example of a trace file from a
+high-quality portion of a sequencing reaction.
+
+Sequence near the beginning or end of a sequencing reaction is often unreliable.
+Although your sequencing results may indicate bases at specific locations, by
+looking at the trace file, you will see that these base calls are unreliable.
+This is an example of a trace file from a portion of a sequencing reaction with
+high background.
+
+If you obtain a different sequence than expected and wish to contact Addgene about
+the accuracy of your plasmid, please email [email protected] and include:
+• The sequencing result
+• The original trace file (.ab1 file) provided by your sequencing company or
+  core facility
+• The corresponding primer sequence used for each reaction
+
+
+TIPS AND FAQs
+================================================================================
+
+My sequence doesn't match Addgene's sequencing result, what should I do?
+
+Check your trace file first; the apparent mismatch/mutation may be the result of a
+mis-called peak in the trace file. If the mutation is not an artifact, please
+email [email protected] with the sequence, trace file, and primer used.
+
+What program can I use to view my trace file?
+
+There are many free programs available that can open .ab1 files, such as:
+• 4Peaks (Mac)
+• SnapGene Viewer (Mac/PC)
+• FinchTV (Mac/PC)
+• Sequence Scanner (PC)
+• Chromas (PC)
+
+How can I tell if a peak is mis-called?
+
+Open the trace file and use the search feature in the program to locate the
+incorrect sequence. Look at the peaks in the area and make sure they are
+justifiable peaks. For instance, in the trace file below, you can see that just
+after base 70 there are multiple peaks in the same location. Looking at the trace
+file will give you more information than simply looking at the bases provided by
+your sequencing provider.
+
+My sequence has "N"s in it- what does that mean?
+
+"N" is the letter used to denote that the position could be any of the 4 bases.
+Check the trace file and see if you can manually call the correct base at the
+position. Sometimes an "N" is the result of an erroneous insertion by the
+sequencing reaction.

biomni/tool/protocols/addgene/Addgene_ Protocol - How to Run an Agarose Gel.txt

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+Title: Addgene: Protocol - How to Run an Agarose Gel
+URL: https://www.addgene.org/protocols/gel-electrophoresis/
+================================================================================
+
+Protocols
+Agarose Gel Electrophoresis
+Agarose Gel Electrophoresis
+You may also like...
+Restriction Digest of Plasmid DNA
+Purifying DNA from an Agarose Gel
+DNA Ligation
+Introduction
+Gel electrophoresis is the standard lab procedure for separating DNA by size (e.g., length in base pairs) for visualization and purification. Electrophoresis uses an electrical field to move the negatively charged DNA through an agarose gel matrix toward a positive electrode. Shorter DNA fragments migrate through the gel more quickly than longer ones. Thus, you can determine the approximate length of a DNA fragment by running it on an agarose gel alongside a DNA ladder (a collection of DNA fragments of known lengths).
+Last Update: Feb. 20, 2018
+Protocol Video
+Watch the protocol video below to learn how to perform gel electrophoresis.
+Equipment
+Casting tray
+Well combs
+Voltage source
+Gel box
+UV light source
+Microwave
+Reagents
+TAE (
+recipe here
+)
+Agarose
+Ethidum bromide (stock concentration of 10 mg/mL)
+Gel loading buffer, New England Biolabs B7022S
+Procedure
+Pouring a Standard 1% Agarose Gel:
+Measure 1 g of agarose.
+Pro-Tip
+Agarose gels are commonly used in concentrations of 0.7% to 2% depending on the size of bands needed to be separated -
+see FAQs below
+. Simply adjust the mass of agarose in a given volume to make gels of other agarose concentrations (e.g., 2 g of agarose in 100 mL of TAE will make a 2% gel).
+Mix agarose powder with 100 mL 1xTAE in a microwavable flask. See
+TAE Recipe
+.
+Pro-Tip
+TBE can be used instead of TAE, labs usually use one or the other, but there is very little difference between the two.
+Note:
+Make sure to use the same buffer as the one in the gel box (do not mix different buffers and do not use water).
+Microwave for 1-3 min until the agarose is completely dissolved (but do not overboil the solution, as some of the buffer will evaporate and thus alter the final percentage of agarose in the gel. Many people prefer to microwave in pulses, swirling the flask occasionally as the solution heats up.).
+Caution
+HOT! Be careful stirring, eruptive boiling can occur.
+Pro-Tip
+It is a good idea to microwave for 30-45 sec, stop and swirl, and then continue towards a boil. Keep an eye on it the solution has a tendancy to boil over. Placing saran wrap over the top of the flask can help with this, but is not necessary if you pay close attention.
+Let agarose solution cool down to about 50 °C (about when you can comfortably keep your hand on the flask), about 5 mins.
+Optional
+: Add ethidium bromide (EtBr) to a final concentration of approximately 0.2-0.5 μg/mL (usually about 2-3 μl of lab stock solution per 100 mL gel). EtBr binds to the DNA and allows you to visualize the DNA under ultraviolet (UV) light.
+Caution
+EtBr is a known mutagen. Wear a lab coat, eye protection and gloves when working with this chemical.
+Note:
+If you add EtBr to your gel, you will also want to add it to the running buffer when you run the gel. If you do not add EtBr to the gel and running buffer, you will need to soak the gel in EtBr solution and then rinse it in water before you can image the gel.
+Pour the agarose into a gel tray with the well comb in place.
+Pro-Tip
+Pour slowly to avoid bubbles which will disrupt the gel. Any bubbles can be pushed away from the well comb or towards the sides/edges of the gel with a pipette tip.
+Place newly poured gel at 4 °C for 10-15 mins OR let sit at room temperature for 20-30 mins, until it has completely solidified.
+Pro-Tip
+If you are in a hurry, the gel will set more quickly if you place the gel tray at 4 °C earlier so that it is already cold when the gel is poured into it.
+Loading Samples and Running an Agarose Gel:
+Add loading buffer to each of your DNA samples and mix well. Use 5 µl of loading buffer per 25 µl of sample.
+Note:
+Loading buffer serves two purposes: 1) it provides a visible dye that helps with gel loading and allows you to gauge how far the DNA has migrated; 2) it contains a high percentage of glycerol that increases the density of your DNA sample causing it settle to the bottom of the gel well, instead of diffusing in the buffer.
+Once solidified, place the agarose gel into the gel box (electrophoresis unit).
+Fill gel box with 1xTAE (or TBE) until the gel is covered.
+Pro-Tip
+Remember, if you added EtBr to your gel, add some to the buffer as well. EtBr is positively charged and will run the opposite direction from the DNA. So if you run the gel without EtBr in the buffer you will reach a point where the DNA will be in the bottom portion of the gel, but all of the EtBr will be in the top portion and your bands will be differentially intense. If this happens, you can just soak the gel in EtBr solution and rinse with water to even out the staining after the gel has been run, just as you would if you had not added EtBr to the gel in the first place.
+Carefully load a molecular weight ladder into the first lane of the gel.
+Note:
+When loading the sample in the well, maintain positive pressure on the sample to prevent bubbles or buffer from entering the tip. Place the very top of the tip of the pipette into the buffer just above the well. Very slowly and steadily, push the sample out and watch as the sample fills the well. After all of the sample is unloaded, push the pipettor to the second stop and carefully raise the pipette straight out of the buffer.
+Carefully load your samples into the additional wells of the gel.
+Run the gel at 80-150 V until the dye line is approximately 75-80% of the way down the gel. A typical run time is about 1-1.5 hours, depending on the gel concentration and voltage.
+Note:
+Black is negative, red is positive. The DNA is negatively charged and will run towards the positive electrode.
+Always Run to Red.
+Turn OFF power, disconnect the electrodes from the power source, and then carefully remove the gel from the gel box.
+Optional:
+If you did not add EtBr to the gel and buffer, place the gel into a container filled with 100 mL of TAE running buffer and 5 μL of EtBr, place on a rocker for 20-30 mins, replace EtBr solution with water and destain for 5 mins.
+Using any device that has UV light, visualize your DNA fragments. The fragments of DNA are usually referred to as ‘bands’ due to their appearance on the gel.
+Pro-Tip
+If you will be purifying the DNA for later use, use long-wavelength UV and expose for as little time as possible to minimize damage to the DNA.
+Note:
+When using UV light, protect your skin by wearing safety goggles or a face shield, gloves and a lab coat.
+Analyzing Your Gel:
+Using the DNA ladder in the first lane as a guide (the manufacturer's instruction will tell you the size of each band), you can infer the size of the DNA in your sample lanes. For more details on doing diagnostic digests and how to interpret them please see the
+Diagnostic Digest
+page.
+Purifying DNA from Your Gel:
+If you are conducting certain procedures, such as molecular cloning, you will need to purify the DNA away from the agarose gel. For instructions on how to do this, visit the
+Gel Purification
+page.
+Tips and FAQ
+How do you get better resolution of bands?
+A few simple ways to increase the resolution (crispness) of your DNA bands include: a) running the gel at a lower voltage for a longer period of time; b) using a wider/thinner gel comb; or c) loading less DNA into the well. Another method for visualizing very short DNA fragments is polyacrylamide gel electrophoresis (PAGE), which is typically used to separate 5 - 500 bp fragments.
+How do you get better separation of bands?
+If you have similarly sized bands that are running too close together, you can adjust the agarose percentage of the gel to get better separation. A higher percentage agarose gel will help resolve smaller bands from each other, and a lower percentage gel will help separate larger bands.
+10% Rule
+For each sample you want to load on a gel, make 10% more volume than needed because several microliters can be lost in pipetting. For example, if you want to load 1.0 μg in 10 μL, make 1.1 μg in 11 μL.
+Reference Page
+Top
+Index

biomni/tool/protocols/addgene/Addgene_ Protocol - How to Streak a Plate.txt

@@ -0,0 +1,91 @@
+PROTOCOL: How to Streak a Plate
+SOURCE: https://www.addgene.org/protocols/streak-plate/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+If you have a glycerol stock or stab culture of bacteria and need to purify plasmid
+DNA from it, you will want to isolate an individual clonal population (single
+colony) of bacteria from this stock. Using a single colony from a freshly streaked
+agar plate to inoculate a bacterial culture for DNA purification will minimize the
+chance of having a mixture of plasmids in your purified DNA. This protocol explains
+how to isolate a single bacterial colony by streaking it onto an LB agar plate.
+
+Last Update: Feb. 28, 2017
+
+
+EQUIPMENT
+================================================================================
+
+• Sterile toothpicks or wire loop
+• Bunsen burner (or other small flame source)
+• Incubator
+• Marker
+
+
+REAGENTS
+================================================================================
+
+• LB agar plate (with appropriate antibiotic)
+• Bacterial stab
+
+
+PROCEDURE
+================================================================================
+
+1. Obtain an LB agar plate with appropriate antibiotic.
+
+2. Label the bottom of the plate with the plasmid name and the date. It is also a
+   good idea to add the antibiotic resistance and your initials. Labeling within a
+   laboratory setting is important for organization, and it is recommended that you
+   keep a standard labeling system for all your objects/solutions.
+
+3. Sterilize your lab bench by spraying it down with 70% ethanol and wiping it down
+   with a paper towel. Maintain sterility by working near a flame or bunsen burner.
+
+4. Obtain the appropriate bacterial stab or glycerol stock.
+
+5. Using a sterile loop, pipette tip or toothpick, touch the bacteria growing within
+   the punctured area of the stab culture or the top of the glycerol stock.
+
+   💡 PRO TIP: If you use a wire loop you can sterilize it by passing it through a
+   flame, just be sure to allow enough time for the loop to cool before touching it
+   to the bacteria.
+
+6. Gently spread the bacteria over a section of the plate, as shown in the diagram
+   above, to create streak #1.
+
+   💡 PRO TIPS:
+   • Hold your tooth pick at an angle, the way you would hold a pencil, so that you
+     can make a broad stroke. Only touch the surface of the plate, do NOT dig into
+     the agar.
+   • Another very popular technique is to draw in discontinuous lines. Start by
+     streaking a vertical line of bacteria along one edge of the plate. Then streak
+     horizontal lines in another section of the plate, and then diagonal lines in
+     another section of the plate. Make sure that the first line (and only the
+     first) in each new section crosses at least one line of the previous section
+     so that it will contain some bacteria.
+
+7. Using a fresh, sterile toothpick, or freshly sterilized loop, drag through
+   streak #1 and spread the bacteria over a second section of the plate, to create
+   streak #2.
+
+8. Using a third sterile pipette tip, toothpick, or sterilized loop, drag through
+   streak #2 and spread the bacteria over the last section of the plate, to create
+   streak #3.
+
+9. Incubate plate with newly plated bacteria overnight (12-18 hours) at 37 °C.
+
+   💡 PRO TIP: Some plasmids or bacteria need to be grown at 30 °C instead of 37 °C.
+   This is often true for large unstable plasmids, which sometimes recombine at 37
+   °C. Be sure to check this before incubating your plate.
+
+10. In the morning, single colonies should be visible. A single colony should look
+    like a white dot growing on the solid medium. This dot is composed of millions
+    of genetically identical bacteria that arose from a single bacterium. If the
+    bacterial growth is too dense and you do not see single colonies, re-streak
+    onto a new agar plate to obtain single colonies.
+
+11. Once you have single colonies, you can proceed to Recovering Plasmid DNA or use
+    the individual colonies for other experiments.

biomni/tool/protocols/addgene/Addgene_ Transfection for Recombinant Antibodies.txt

@@ -0,0 +1,231 @@
+Title: Addgene: Transfection for Recombinant Antibodies
+URL: https://www.addgene.org/protocols/transfection-for-recombinant-antibodies/
+================================================================================
+
+Protocols
+Transfection for Recombinant Antibodies
+Transfection for Recombinant Antibodies
+You may also like...
+Antibody Plasmid Collection
+Antibodies Blog Posts
+Antibody Guide
+General Transfection Protocol
+Recombinant Antibody Purification Protocol
+Introduction
+Transfections allow for transient expression of a gene of interest in cell culture. This protocol describes how to transfect suspension HEK293 cells with recombinant antibody plasmids using Polyethylenimine Max as a transfection reagent. After transfection and expression, the recombinant antibody can be purified for use in a variety of applications.
+Sharing speeds science.
+We believe that sharing the full details of our protocols supports reproducibility and accelerates science. Here, we list the specific equipment, reagents, and methods that we use in our lab at Addgene. Equipment and reagents from other vendors should produce similar outcomes when using these protocols. However, please be aware that your protocol may need to be adjusted to accommodate slight differences between products. Addgene does not endorse or recommend specific products or equipment. Inclusion of this information is solely for transparency intended to support reproducibility in science.
+General Considerations
+For antibody-expressing plasmids containing the SV40 origin of replication use a HEK293 line stably expressing the SV40 large T antigen.
+For antibody-expressing plasmids containing the Epstein-Barr virus origin of replication use a HEK293 line stable expressing Epstein Barr nuclear antigen (EBNA).
+Safety Warnings
+HEK293 cells are considered biosafety level 2. Please ensure that you are in compliance with your institution’s biosafety regulations.
+Last Update: February 18, 2022
+Workflow Timeline
+Day 1:
+Seed cells
+Day 2:
+Transfect cells
+Day 3-6:
+Feed cells
+Day 7:
+Harvest antibody
+Equipment
+Biosafety Cabinet
+4 °C Refrigerator
+Micropipettes and tips
+Pipettor and pipettes
+Benchtop centrifuge compatible with 50 mL conical tubes
+Automated cell counter
+37 °C, 5% CO
+2
+incubator with shaking platform set to 120 rpm
+37 °C bead bath
+Vortex
+Stir bar
+Magnetic stir plate
+pH meter
+Reagents
+HEK293 cells
+Recombinant antibody plasmid DNA
+500 mL vented flask
+50 mL conical tubes
+Microcentrifuge tubes
+Cell counting chamber slide
+30 mL luer-lock syringe, BD BD302832
+0.2 µm luer-lock filter, VWR 431229
+0.22 µm PES filtering system, 1000 mL, VWR 431098
+0.45 µm PES filtering system, 500 mL, VWR 430770
+Trypan Blue, Thermo Fisher T10282
+Valproic acid sodium salt, Sigma Aldrich P4543-10G
+Polyethylenimine hydrochloride, M.W. 40000 (PEI-MAX), Linear, Transfection Grade, VWR 75800-188
+10% Poloxamer 188, Thermo Fisher 24040032
+Glutagro, Corning 25-015-CI
+BalanCD HEK293 Media, Irvine Scientific 91165-1L
+BalanCD HEK293 Feed, Irvine Scientific 91166-500ML
+Benzamide, Millipore Sigma 12072
+Antipain, Millipore Sigma 10791
+Leupeptin, Millipore Sigma L2884
+Aprotinin saline solution, Millipore Sigma A6279
+Before Starting
+Warm the DNA and working stock of PEI-MAX to room temperature before use, and warm the BalanCD HEK293 transfection media (BCD TFX) to 37 °C. See below for other Reagent Preparation instructions.
+Wipe down all pipettes and equipment with 10% bleach prior to use.
+Reagent Preparation
+100 mM Valproic Acid
+Dissolve
+2.88 g
+valproic acid in
+200 mL
+deionized water
+Sterilize by passage through a sterile 0.22 µm filter.
+Prepare 10 mL aliquots and store the solution at -20 °C.
+1 mg/mL PEI-MAX
+Add
+1 g
+of PEI-MAX powder to
+900 mL
+deionized water in a 1 L bottle and stir on a magnetic stir plate.
+Stir until all particles have dissolved. Note: The powder should dissolve rapidly (with 20 min) but check for the presence of particles still in solution. Continue to stir until all particles have dissolved. This may take several hours.
+Adjust to
+pH 7.0
+with 10 N sodium hydroxide (NaOH) or 5 N hydrochloric acid (HCl). Note: Adjust the pH slowly, adding no more than 200 µL of NaOH or 20 µL of HCl at a time.
+Add deionized water to a final volume of
+1 L
+and recheck pH to ensure that it has not drifted.
+Filter the solution through a 0.22 µm filter.
+Prepare aliquots and store at -20 °C until use.
+BCD TFX
+1000 mL
+BalanCD HEK293 Media
+10 mL
+10% Poloxamer 188
+40 mL
+200 mM Glutagro
+Do not add selective reagents
+Store at 4 °C until use. We suggest preparing fresh solutions after one month.
+BCD Feed
+500 mL
+BalanCD HEK293 Feed
+20 mL
+200 mM Glutagro
+Store at 4 °C until use. We suggest preparing fresh solutions after one month.
+1000X protease inhibitor cocktail
+25 mg
+Leupeptin
+50 mg
+Antipain
+250 mg
+Benzamidine
+25 mL
+Aprotinin saline solution (2 mg/mL)
+Mix well and sterilize through a 0.2 µm PES syringe filter.
+Aliquot and freeze upright at -20 °C.
+Procedure
+Section 1: Seeding cells
+The day prior to transfecting, seed a
+108 mL
+culture of HEK293 cells at a density of
+0.9 x 10
+6
+cells/mL
+in a 500 mL vented flask.Incubate in a
+37 °C, 5% CO
+2
+incubator on a shaking platform set to 120 rpm.
+Pro-Tip
+Do not use cells that are over 30 passages.
+Section 2: Transfection
+Check the cell density and viability:
+Using a 5 mL pipette, transfer
+0.5 mL
+of HEK293 cell suspension into a clean microcentrifuge tube.
+Pro-Tip
+Cells settle quickly and need to be resuspended before sampling. Gently swirl the flask 5–10 times before sampling.
+Transfer
+10 µL
+of trypan blue into a clean microcentrifuge tube.
+Vortex the cell suspension.
+Transfer
+10 µL
+of cell suspension into the microfuge tube containing the trypan blue.
+Pipette 10 times to mix.
+Load
+10 µL
+of the cell suspension/trypan blue mix into a cell counting chamber.
+Load the cell counting chamber on an automated cell counter and measure the live cell density and viability of the culture.
+Pro-Tip
+Culture should be between
+1.5–2 x 10
+6
+cells/mL
+with
+>95%
+viability to proceed with transfection.
+Transfect the flask containing 108 mL cells:
+Transfer
+6 mL
+of BCD TFX into each of two 50 mL tubes.
+Cap the tubes and incubate for
+1 h
+in the
+37 °C
+bead bath.
+Transfer the PEI-MAX and recombinant antibody plasmid DNA sample to the biosafety cabinet and incubate for
+1 h
+at room temperature.
+After the BCD TFX has warmed in the bead bath, transfer the tubes to the biosafety cabinet.
+Add
+180 µg
+of recombinant antibody plasmid DNA to one tube of 6 mL BCD TFX. Cap the tube and vortex for 5 s to mix.
+Add
+450 µL
+of 1 mg/mL PEI-MAX to the second tube of 6 mL BCD TFX (for a 1 mg/mL stock solution of PEI-MAX, 450 µg = 450 µL). Cap the tube and vortex for 5 s to mix.
+Pro-Tip
+The optimal ratio of DNA:PEI may vary significantly and should be empirically determined for your sample. Typical ratios may range from 1:1 to 1:6.
+Add the diluted PEI-MAX to the diluted DNA. Cap the tube and vortex with three 1 s pulses.
+Incubate for
+3 min
+at room temperature.
+Transfer the flask of HEK293 cells to the biosafety cabinet.
+Add
+12 mL
+of PEI-MAX/DNA mix to the flask dropwise.
+Cap the flasks and swirl 5–10 times to mix.
+Return the flask to the incubator.
+Section 3: BCD Feed and valproic acid supplementation
+During the
+24–144 h
+post-transfection, supplement the flask with 4% BCD Feed.
+Pro-Tip
+The feed can be repeated up to 4 times for a total of 16% of the culture volume.
+At
+72-96 h
+post-transfection, supplement the flask with
+3.75 mM
+valproic acid.
+Example feeding schedule:
+Thursday: Transfect cells.
+Friday (24 h post-transfection): Add 4% BCD Feed.
+Monday (96 h post-transfection): Add valproic acid to 3.75 mM, add 4% BCD Feed.
+Tuesday (120 h post-transfection): Add 4% BCD Feed.
+Wednesday (144 h post-transfection): Add 4% BCD Feed.
+Thursday (168 h post-transfection): Harvest antibody.
+Section 4: Harvest antibody
+At 168 hours (1 week) post-transfection, harvest the antibody.
+Transfer the HEK293 cells and media to 50 mL conical tubes.
+Centrifuge for
+15 min
+at
+3100 x
+g
+to pellet the cells.
+Filter the supernatant through a 0.45 µm PES filter.
+Add
+1X
+protease inhibitor cocktail to the supernatant.
+The supernatant containing the recombinant antibody can be used as-is or the recombinant antibody can be purified from the supernatant (see our
+Recombinant Antibody Purification Protocol
+).
+Store the supernatant at
+4 °C
+until ready to use.

biomni/tool/protocols/addgene/Addgene_ Using a Light Microscope Protocol.txt

@@ -0,0 +1,151 @@
+PROTOCOL: Using a Light Microscope Protocol
+SOURCE: https://www.addgene.org/protocols/light-microscope/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+Microscopes are emblematic of biological research and are found in many different
+types of laboratories. These tools allow you to observe specimens much smaller than
+you would be able to with your naked eye, like microbes in a drop of pond water or
+the cells in your cell culture dish. Microscopes come in a huge range of shapes and
+sizes - from phone-sized, foldable microscopes that aren't very powerful (but are
+cheap and accessible) to massive transmission electron microscopes that allow us to
+see cellular components with science-fiction levels of detail. You can take entire
+courses on microscopy and still have more to learn, so, for this protocol, we'll
+focus on one of the most common types that you're likely to encounter: the compound
+light microscope.
+
+As the name suggests, light microscopes take advantage of the physical properties of
+light to detect small objects. Two of the most important properties of a microscope
+are magnification (the ability to make an image larger) and resolution (the ability
+to distinguish between two discrete objects). Figure 1 depicts an image of a compound
+light microscope with the main components labeled:
+
+Main Components:
+• Base
+• Light Source
+• Condenser and Diaphragm
+• Stage
+• Objective Lenses
+• Focus Knobs (Fine and Coarse)
+• Nosepiece
+• Arm
+• Ocular Lens (Eyepiece)
+
+The route the light follows from the source to your eyes is called the light path.
+Light travels from the light source through a condenser that helps concentrate the
+light onto the sample, and then through the sample. However, some of that light
+won't make it through the sample because different components of the specimen will
+refract and reflect the light. These differences create contrast, which allows you
+to distinguish objects within the sample. You can increase contrast by a) adjusting
+the aperture of the condenser diaphragm to limit the amount of light hitting the
+sample or b) using dyes or stains that add color to some components of the sample
+but not others.
+
+The light that does pass through the sample then travels through the objective lens
+which magnifies the image, then the ocular lens, where the image is further
+magnified before it finally reaches your eyes. These lenses determine the
+magnification of the image and the resolution your microscope can achieve. You
+achieve the highest resolution when your image is in focus, which you can control
+using the focus knobs on the side of the microscope.
+
+You may be wondering why there are multiple objective lenses. Each objective performs
+essentially the same task but has different magnifying powers. When you look at the
+side of an objective, you will see several pieces of information, but the most
+important (for now) is the magnification power, such as 4x, 10x, or 20x. A 4x
+objective magnifies the image four times the actual size, the 10x objective
+magnifies it 10 times, and so on.
+
+The ocular lens also provides magnification and the power should be provided on the
+microscope; often this lens provides 10x magnification. To determine the final
+magnification of your image, multiply the magnification of your objective lens by
+the magnification of your ocular lens. For example, if you observe something using
+the 10x objective and your ocular lens is also 10x, you would actually be seeing a
+100x magnified image of that object!
+
+
+EQUIPMENT
+================================================================================
+
+• Light Microscope
+• Prepared Slides (or other sample that can fit on a microscope stage)
+
+
+REAGENTS
+================================================================================
+
+None needed
+
+
+PROCEDURE
+================================================================================
+
+1. Set up your microscope by removing any protective cover that may be on it and
+   plugging in the power source for the light.
+
+   💡 PRO TIP: If you have to move your microscope around the lab, be sure to
+   carry it with two hands - one hand supporting the base and the other holding
+   the arm - or use a cart.
+
+2. Revolve the nosepiece so that the lowest power objective is directly over the
+   stage. It is best practice to start with the lowest power objective to find your
+   sample.
+
+3. Place your slide (or other sample type) on the microscope stage. If using a
+   slide, you can secure it into place using the metal clips on the stage.
+
+4. Turn on the power source and use the stage arm to move the stage so that the
+   light shines onto your sample.
+
+5. To ensure that your sample is adequately illuminated, look through the
+   eyepiece. Adjust the power of the light source so that you can look through the
+   eyepiece comfortably. Then, close down the diaphragm and use the coarse focus
+   adjustment knob (the larger of the two focus knobs) until you can see the edges
+   of the aperture are in focus. Center the aperture in your field of view and
+   then slowly open the diaphragm until the edges are just out of view.
+
+   💡 PRO TIP: Looking through a microscope's eyepiece can take some practice.
+   Try to keep both eyes open and adjust the position of the eyepiece so that you
+   can see your image through both sides.
+
+6. Once you are happy with the lighting, use the coarse focus knob to bring your
+   sample into focus. Once your sample is in focus, use the stage arm to move the
+   stage (and ultimately your slide) until your field of view is centered on a
+   region of interest. If needed, you can also use the fine focus knob (the smaller
+   of the focus knobs) to make minor adjustments to the focus.
+
+7. After you are happy with the positioning and focus of your image on the lowest
+   magnification, you are ready to move up to the next objective. Take your eyes
+   away from the eyepiece and carefully revolve the nosepiece so that the next
+   objective is in position over your sample. As you increase magnification, you
+   will notice that the objectives get larger. Be careful to avoid hitting your
+   sample with the objective!
+
+8. Once on the higher objective, you will likely need to re-adjust lighting and
+   focus. Increase the power of the light source by opening the diaphragm so that
+   you can easily see your sample. Then use the focus knobs to bring the sample
+   into focus.
+
+   💡 PRO TIP: As you increase your magnification, remember that the objectives
+   are larger. Adjust your focus slowly so that you don't accidentally hit your
+   slide and risk damaging the lens or your sample.
+
+9. You can repeat steps 7 and 8 with progressively higher magnification lenses.
+
+10. Depending on why you are using the microscope you may wish to record your
+    observations. Some microscopes have digital cameras attached to them that
+    allow you to capture images directly to a computer, but many microscopes do
+    not and you may have to dust off your drawing skills. Regardless of your image
+    capture method, be sure to record the magnification used, any stains used in
+    your sample, and sample information so that you or others can interpret your
+    images in the future.
+
+
+CONCLUSION
+================================================================================
+
+Like any important laboratory instrument, you should be sure to take care of your
+microscope. Bumps, scratches, and dust can easily impair microscope performance. Be
+sure to clean up your work area and microscope after use and store the microscope
+appropriately.

biomni/tool/protocols/addgene/Addgene_ Western Blot.txt

@@ -0,0 +1,308 @@
+PROTOCOL: Western Blot
+SOURCE: https://www.addgene.org/protocols/western-blot/
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+Western blot is a technique used to separate proteins by size followed by
+detection using antibodies specific to the protein of interest. This protocol
+describes the basic steps for lysing cells, determining total protein
+concentration in the lysate, running a precast SDS-PAGE gel, and immunoblotting.
+
+General Guidelines
+
+• Acrylamide percentage varies in SDS-PAGE gels. Use lower percentage acrylamide
+  when immunoblotting high molecular weight proteins and higher percentage
+  acrylamide when immunoblotting low molecular weight proteins.
+
+• This protocol uses a dry transfer device but can be adapted for wet and
+  semi-dry transfer approaches.
+
+Last Update: January 24, 2022
+
+
+WORKFLOW TIMELINE
+================================================================================
+
+Day 1: Prepare lysates, run SDS-PAGE, transfer, block, incubate with primary
+       antibody
+
+Day 2: Incubate with secondary antibody
+
+
+EQUIPMENT
+================================================================================
+
+• Microcentrifuge
+• 0.5–10 µL single channel pipette
+• 2–20 µL single channel pipette
+• 20–200 µL single channel pipette
+• 200–1000 µL single channel pipette
+• Pipette controller
+• Pipette tips and pipettes
+• Spectrophotometer
+• Heat block
+• Mini gel tank chamber
+• Power supply
+• iBlot 2 Gel Transfer Device
+• Roller
+• Spatula
+• Platform shaker
+• Cold room
+• Gel imager
+• -80 °C freezer
+
+
+REAGENTS
+================================================================================
+
+• 1X PBS
+• Lysis buffer e.g., RIPA lysis buffer
+• Microcentrifuge tubes
+• BCA Assay, Thermo Fisher 23227
+• β-mercaptoethanol
+• 4X protein loading buffer
+• Precast SDS-PAGE gel
+• SDS-PAGE running buffer
+• Prestained protein ladder
+• Ethanol
+• iBlot 2 PVDF Mini Stack, Thermo Fisher IB24002
+• 20X TBS
+• Tween-20
+• Nonfat milk powder
+• 96-well microtiter plate
+• Chemiluminescence substrate
+• Plastic wrap
+• Primary antibody
+• Secondary antibody
+• Deionized water
+
+
+BEFORE STARTING
+================================================================================
+
+• Refer to the manufacturer's instructions for additional information specific to
+  your antibody, such as ideal blocking buffer and optimal antibody
+  concentrations. Consider titrating your antibody to determine the optimal dose.
+
+• Secondary antibodies must match the host species of the primary antibody. For
+  example, use an anti-mouse secondary antibody for primary antibodies raised in
+  a mouse.
+
+
+PROCEDURE
+================================================================================
+
+Section 1: Lyse cells
+
+1. Centrifuge 5 x 10⁶ cells for 5 min at 100 x g.
+
+2. Gently remove supernatant.
+
+3. Resuspend cell pellet in 1 mL of 1X PBS and transfer to a microcentrifuge
+   tube.
+
+4. Centrifuge for 5 min at 100 x g.
+
+5. Carefully remove supernatant.
+
+6. Resuspend cell pellet in appropriate volume of cold lysis buffer.
+
+   💡 PRO TIP: The volume of lysis buffer will vary depending on the size of your
+   cell pellet but will generally be between 250–1000 µL.
+
+   💡 PRO TIP: The ideal lysis buffer will vary depending on the cellular location
+   of the protein of interest. RIPA buffer is suitable for most proteins but more
+   stringent buffers and a sonication step may be required for hard to extract
+   proteins such as those in the nucleus.
+
+7. Incubate on ice for 30 min.
+
+8. Centrifuge lysate for 15 min at 14,000 x g at 4 °C.
+
+9. Transfer supernatant to a clean microcentrifuge tube.
+
+10. Use the lysate immediately or store at -80 °C until ready to use.
+
+Section 2: Determine the total protein concentration and prepare the lysate for
+SDS-PAGE
+
+1. Determine the protein concentration using a Pierce BCA Assay Kit or other
+   preferred method for protein determination.
+
+2. Prepare a 50:1 Reagent A to Reagent B dilution of the BCA assay.
+
+3. Prepare serial dilutions of the BSA standard that range from 0–2000 µg/mL.
+
+4. In duplicate, dilute 10 µL of standard, blank, and lysate samples into 200 µL
+   of BCA reagent in a 96-well microtiter plate.
+
+5. Incubate for 30 min at 37 °C.
+
+6. Determine the absorbance at 590 nm.
+
+7. Calculate the average absorbance of the duplicate samples on the plate.
+
+8. Subtract the average absorbance of the blank from all of the samples.
+
+9. Plot a standard curve of BSA standard concentration versus absorbance.
+
+10. Extrapolate the total protein concentration of the sample from the standard
+    curve.
+
+11. Determine the volume of sample required to load an equivalent amount of total
+    protein for each sample.
+
+    💡 PRO TIP: The ideal total protein loaded will vary between samples and
+    target proteins but is typically between 10–50 µg. If the protein is in low
+    abundance in the sample you will need to load a greater amount of total
+    protein.
+
+12. Prepare sample for loading as follows:
+    • Add 10% v/v β-mercaptoethanol to the 4X protein loading buffer.
+    • Dilute 4X protein loading buffer in the sample to 1X.
+    • Boil the samples for 10 min at 100 °C.
+
+Section 3: SDS-PAGE
+
+1. Prepare the precast gel as follows:
+   • Remove the gel from the plastic packaging.
+   • Remove tape and plastic comb.
+   • Rinse the wells with deionized water 3x.
+   • Shake the gel gently between washes to remove residual water.
+
+2. Load the gel into the chamber of the SDS-PAGE gel tank.
+
+   💡 PRO TIP: Different SDS-PAGE gel tanks will have different gel orientations.
+   Refer to the instructions for your specific tank.
+
+3. Close the clamp.
+
+4. Prepare 1X running buffer as follows:
+   • Dilute 25 mL of 20X running buffer to 500 mL with deionized water. Mix well.
+
+5. Fill the chamber of the SDS-PAGE gel tank with the 1X running buffer.
+
+6. Load the samples into the gel.
+
+7. Load 5–10 µL of the prestained protein ladder.
+
+8. Place the lid on the tank and plug the electrode cords into the power supply.
+
+9. Run the gel at 100 V for 10–15 min or until the samples have moved out of the
+   wells and into the gel.
+
+10. Increase the voltage to 150 V and continue running the gel until you have
+    obtained the desired separation.
+
+11. Gently open the gel with the spatula.
+
+Section 4: Dry transfer
+
+1. Soak the gel for 15 min in 20% ethanol in deionized water.
+
+   To prepare 20% ethanol, dilute 2 mL of ethanol into 8 mL of deionized water
+   and mix well.
+
+2. Assemble the transfer sandwich as follows:
+   • Unseal the iBlot 2 PVDF Mini transfer stack.
+   • Set the Top Stack to one side and discard the white separator.
+   • Keep the Bottom Stack in the plastic tray.
+   • Place the Bottom Stack on the blotting surface.
+   • Align the electrical contacts on the blotting surface of the iBlot 2 Gel
+     Transfer Device.
+   • Wet the pre-run gel in deionized water and place it on the transfer membrane
+     of the bottom stack.
+   • Soak a piece of iBlot Filter Paper in deionized water.
+   • Place the pre-soaked iBlot Filter Paper on the gel and remove the air bubbles
+     with the roller.
+   • Place the Top Stack over the pre-soaked filter paper.
+   • Remove air bubbles using the roller.
+   • Place the Absorbent Pad on top of the Top Stack such that the electrical
+     contacts are aligned with the corresponding electrical contacts on the
+     blotting surface of the iBlot 2 Gel Transfer Device.
+
+3. Close the lid of the device.
+
+4. Select the desired method and make sure the parameters are correct.
+
+5. Transfer for 5–6 min for proteins <30 kDa.
+
+6. Transfer for 8–10 min for proteins >150 kDa.
+
+7. Select Start Run.
+
+8. When the run is complete, select Done.
+
+Section 5: Blocking
+
+1. Prepare 1X TBST as follows:
+   • 25 mL of 20X TBS
+   • 2.5 mL of Tween-20
+   • 472.5 mL of deionized water
+   • Mix well
+
+2. Prepare blocking buffer as follows:
+   • Dilute 5% w/v non-fat milk into 1X TBST
+   • Example: 5 g of non-fat milk into 100 mL of 1X TBST.
+
+   💡 PRO TIP: The ideal blocking buffer will vary between antibodies. Refer to
+   the manufacturer's information before running your experiment.
+
+3. Remove the membrane from the transfer device and place it in a container with
+   1X TBST, protein side up.
+
+4. Block the membrane in blocking buffer for 1 h at RT on a shaking platform.
+
+5. Wash the membrane 3x for 5 min in 1X TBST at RT on a shaking platform.
+
+Section 6: Antibody incubation
+
+1. Dilute the primary antibody to the desired concentration in blocking buffer.
+
+   💡 PRO TIP: The optimal concentration will vary between antibodies but is
+   typically between 1–10 μg/mL.
+
+2. Incubate the membrane overnight in primary antibody at 4 °C on a rocking
+   platform.
+
+   💡 PRO TIP: Primary antibody incubation can be reduced to 2 h at RT but may
+   result in more non-specific binding.
+
+3. Wash the membrane 3x for 5 min in 1X TBST at RT on a shaking platform.
+
+4. Dilute the horseradish peroxidase-conjugated secondary antibody to the desired
+   concentration in blocking buffer.
+
+   💡 PRO TIP: The optimal concentration will vary between antibodies but is
+   typically between 1–10 μg/mL.
+
+5. Incubate the membrane with secondary antibody for 60 min at RT on a shaking
+   platform.
+
+6. Wash the membrane 3x for 5 min in 1X TBST at RT on a shaking platform.
+
+7. Prepare the chemiluminescence substrate by mixing 1:1 reagent A to reagent B.
+
+8. Gently incubate the membrane in the chemiluminescence reagent for 5 min at RT.
+
+9. Cover the membrane with clear plastic and use a gel imager with
+   chemiluminescence detection or a dark room to detect the bands.
+
+
+TIPS AND TROUBLESHOOTING
+================================================================================
+
+• The optimal lysis buffer will vary depending on the sample type and cellular
+  location of the protein of interest. You may need to try a variety of lysis
+  buffers to find the best for your target.
+
+• The optimal concentration and blocking buffer will vary between antibodies.
+  Review the instructions before starting your experiment and consider titrating
+  your antibody to determine the optimal dose.
+
+• To ensure that your antibody is both functioning as expected and is specific,
+  include a positive control sample that you know expresses the protein, such as
+  cells transfected with the protein of interest, and a negative control sample
+  that does not express the protein of interest.

biomni/tool/protocols/addgene/Addgene_ What is Polymerase Chain Reaction (PCR).txt

@@ -0,0 +1,226 @@
+PROTOCOL: What is Polymerase Chain Reaction (PCR)
+SOURCE: https://www.addgene.org/protocols/pcr/
+================================================================================
+
+BACKGROUND INFORMATION
+================================================================================
+
+A standard Polymerase Chain Reaction (PCR) is an in vitro method that allows a
+single, short region of a DNA molecule (single gene perhaps) to be copied multiple
+times by Taq Polymerase. From a single copy of DNA (the template), a researcher can
+create thousands of identical copies using a simple set of reagents and a basic
+heating and cooling (denaturing and annealing) cycle. The process became automated
+with the discovery of a heat resistant DNA polymerase from the thermophilic
+bacterium, Thermus aquaticus (Taq). Taq polymerase can withstand many heating and
+cooling cycles, which would denature DNA polymerases from other species.
+
+In addition to the template DNA and the Taq polymerase, PCR requires free
+nucleotides [dNTPs; adenine (A), cytosine (C), guanine (G), thymine (T)] in an
+equal molar ratio. It also requires two unique single stranded DNA oligonucleotide
+(oligo) primers, which anneal to the regions upstream (5') and downstream (3') of
+the DNA segment to be amplified. When these reagents are combined in an appropriate
+buffer, a series of heating (denaturing) and cooling (annealing) steps allow the
+Taq polymerase to copy the DNA in between the oligo primers. This molecular biology
+technique creates several micrograms of target DNA from just a few nanograms of
+template DNA through several cycles of denaturation, annealing, and synthesis. After
+the PCR is complete, the product can be verified based on size by gel
+electrophoresis.
+
+
+BASIC PCR PROGRAM
+================================================================================
+
+1. Initial Denaturation for 2 minutes at 94°C:
+   This initiation step heats the double stranded DNA template strand to the point
+   where the strands start denaturing and the hydrogen bonds are broken between the
+   nucleotide base pairs. The initial denaturation step is commonly performed at
+   94–98°C.
+
+2. Denature 30 seconds at 94°C:
+   Continued denaturation of double stranded DNA.
+
+3. Anneal primers for 30 seconds at 55°C:
+   The forward and reverse primers are stable within this temperature range to
+   anneal to each of the single stranded DNA template strands. The DNA polymerase
+   is also stable enough to now bind to the primer DNA sequence.
+
+4. Extend DNA for 1 minute at 72°C:
+   The Taq polymerase has an optimal temperature around 70-75°C so this step enables
+   the DNA polymerase to synthesize and elongate the new target DNA strand
+   accurately and rapidly.
+
+5. Repeat steps 2-4 25-30 times.
+
+6. Final Extension for 5 minutes at 72°C:
+   A final extension to fill-in any protruding ends of the newly synthesized
+   strands.
+
+
+MATERIALS LIST
+================================================================================
+
+Reagents for each 50µL PCR:
+
+• Thin-walled PCR tube
+• Ice Bucket
+• 2 μL Template DNA (10 ng-500 ng)
+• 5 μl 10X Taq buffer with MgCl₂
+• 1 μl dNTP mix (10 mM each nt)
+• 2.5 μL Forward Primer (10 μM stock)
+• 2.5 μL Reverse Primer (10 μM stock)
+• 36.8 μL Sterile dH₂O
+• 0.2 μL Taq DNA Polymerase (5 units/μL)
+• PCR Machine
+• Agarose Gel
+
+
+PROCEDURE
+================================================================================
+
+Primer Design and PCR
+
+1. Design Primers. See our protocol on how to design primers.
+
+   Note: Primer3 is an excellent resource for choosing primers.
+
+2. Place thin-walled PCR tubes on ice.
+
+3. Set up a 50 μL reaction (Keep all your reagents on ice):
+   • 2 μL Template DNA (10 ng-500 ng)
+   • 5 μl 10X Taq buffer with MgCl₂
+   • 1 μl dNTP mix (10 mM each nt)
+   • 2.5 μL Forward Primer (10 μM stock)
+   • 2.5 μL Reverse Primer (10 μM stock)
+   • 0.2 μL Taq DNA Polymerase (5 units/μL)
+   • 36.8 μL Sterile dH₂O (variable)
+
+   Note: If you are doing multiple PCRs, save time by creating a "master mix,"
+   which minimizes the number of small volumes you need to pipet. If you are using
+   the same template DNA for all your reactions, the Template DNA can be added to
+   the master mix. Forward and Reverse Primers DO NOT get added to a master mix.
+
+4. Place reaction tubes in PCR machine.
+
+5. Set annealing temperature 5°C below the primer melting temperature (Tm).
+
+6. Set extension step at 1-2 minutes per kilobase of product depending on whether
+   you are using a polymerase with proofreading capabilities.
+
+   Note: See manufacturer's instructions for specific instructions about extension
+   time and temperatures.
+
+7. Initial Denaturation for 2 minutes at 94°C.
+
+8. Denature for 30 seconds at 94°C.
+
+9. Anneal primers for 30 seconds at 55°C (or 5°C below Tm).
+
+10. Extend DNA for 2 minutes at 72°C.
+
+11. Repeat steps 8-10 for 25-30 cycles.
+
+12. Final Extension for 5 minutes at 72°C.
+
+13. Run 2 μL on a gel to check size and concentration of PCR product.
+
+Master Mix Preparation
+
+1. Multiply the volume of each reagent by the number of individual PCRs you wish to
+   perform and add 10% extra to account for pipetting error. In this example, we
+   have 7 different reactions (7 unique primer pairs), so we multiply each volume
+   by 7.
+
+2. In a single 1.5mL tube combine the following:
+   • 10X Taq buffer with MgCl₂: 5μl x 7 reactions = 35μl total + 10% (3.5μl) =
+     38.5μl
+   • dNTP mix (10 mM each nt): 1µl x 7 reactions = 7µl total + 10% (0.7µl) =
+     7.7µl
+   • Template DNA: 2µl x 7 reactions = 14µl total + 10% (1.4µl) = 15.4µl
+   • Sterile dH₂O: 36.8µl x 7 reactions = 257.6µl + 10% (25.7µl) = 283.3µl
+   • Taq DNA polymerase: 0.2µl x 7 reactions = 1.4µl + 10% (0.14µl) = 1.54µl
+
+3. Mix the contents by gently pipetting up and down several times. Keep tube on ice.
+
+4. Add the forward and reverse primers to the thinned walled PCR tubes.
+
+   Note: Do this before adding the master mix so you know that the primers have
+   been added, pipet the forward primer onto one side of the tube wall and the
+   reverse primer onto the other.
+
+5. Add the master mix to the thin walled PCR tubes. Put 50µl – 2.5µl (fwd primer) –
+   2.5µl (rev primer) = 45µl volume of master mix to add to each PCR tube.
+
+6. Secure the tops to the PCR tubes, gently tap each tube to bring all the liquid
+   to the bottom before placing it in the PCR machine.
+
+Diluting Primers
+
+Most people order primers from a company, which synthesizes and ships them as a
+lyophilized powder. The researcher then needs to reconstitute their primers in
+liquid, normally sterile dH₂O. To make a 100uM stock of any primer, add a number of
+µl of dH₂O equal to the number of nanomoles of DNA times 10. For example, if your
+lyophilized DNA is 38.5nm, add 385µl of water.
+
+After making your 100uM stock, immediately make a working concentration of each
+primer (10uM) by making a 1:10 dilution of the stock. For example, add 100µl of
+primer stock to 900µl of dH₂O.
+
+
+TIPS AND FAQ
+================================================================================
+
+Tips
+
+• If your target DNA sequence is GC-rich, increase the time of the denaturing.
+
+• Your 5' and 3' primers should be designed to have similar melting temperatures
+  (Tm). Set the annealing temperature to 5°C lower than the Tm of your primers. If
+  you are getting non-specific PCR products, increase the annealing temperature
+  step-wise by 1-2°C.
+
+• The rate of DNA synthesis is ~1-2 kb/min. The extension time can be adjusted
+  according to the length of the target sequence.
+
+FAQ
+
+How do I design primers?
+
+See our reference page on how to design primers for detailed instructions and
+check out our protocol on plasmid cloning by PCR to learn how to design primers for
+cloning purposes.
+
+What do I do if my PCR isn't working?
+
+Try adding 1µl of 25mM MgCl₂ and/or 1µl DMSO to each reaction. The ideal setup for
+this troubleshooting step is to do one reaction with each, and one reaction with
+both. MgCl₂ acts to supplement the amount supplied by the buffer stock, where it is
+known to form a gradient when frozen. DMSO helps with denaturation of the DNA,
+particularly for GC-rich templates.
+
+What does each ingredient specifically do?
+
+• Template DNA: Contains the portion of DNA that we are interested in amplifying
+  for analysis and manipulation. It is essential that the template DNA and primer
+  are complementary for a PCR to work efficiently.
+
+• Taq Buffer with MgCl₂: Provides an optimal and stable chemical environment for
+  the DNA polymerase to work adequately. Divalent cations such as Mg²⁺ and Mn²⁺
+  stabilize the buffer solution. These cations can also be used for PCR-mediated DNA
+  mutagenesis. A higher cation concentration increases the error rate of the DNA
+  polymerase.
+
+• Deoxynucleoside Triphosphates (dNTPs): Are the building blocks added one at a time
+  to the new DNA strand by the DNA polymerase.
+
+• Forward and Reverse Primers: Hybridize and are complementary to the 3' ends of
+  the sense and anti-sense strands of the template DNA. They are strands of nucleic
+  acid that are starting points for DNA elongation and synthesis.
+
+• Taq DNA polymerase: Special DNA polymerase that can withstand radical
+  temperature changes during a typical PCR. The DNA polymerase has an optimum
+  temperature around 70°C and is the molecule responsible for driving the DNA
+  synthesis.
+
+• Sterile dH₂O: Used to fill the remaining 50 μL of the reaction mix. Its solvent
+  and buffer capabilities provide a suitable environment for the DNA amplification
+  reaction.

biomni/tool/protocols/thermofisher/Actin Staining Protocol _ Thermo Fisher Scientific - US.txt

@@ -0,0 +1,229 @@
+PROTOCOL: Actin Staining Protocol
+SOURCE: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/microscopy-protocol/actin-staining-protocol.html
+================================================================================
+
+INTRODUCTION
+================================================================================
+
+We offer several fluorescent and biotinylated phalloidin and phallacidin derivatives
+for labeling F-actin. These phallotoxins, isolated from the deadly Amanita phalloides
+mushroom, are bicyclic peptides that differ by two amino acid residues. They can be
+used interchangeably in most applications and bind competitively to the same sites in
+F-actin. Phalloidin and phallacidin contain an unusual thioether bridge between a
+cysteine and tryptophan residue that forms an inner ring structure. At elevated pH,
+this thioether is cleaved and the toxin loses its affinity for actin.
+
+Fluorescent and biotinylated phallotoxins stain F-actin at nanomolar concentrations
+and are extremely water soluble, thus providing convenient probes for labeling,
+identifying, and quantitating F-actin in tissue sections, cell cultures, or cell-free
+experiments. Labeled phallotoxins have similar affinity for both large and small
+filaments, binding in a stoichiometric ratio of about one phallotoxin molecule per
+actin subunit in muscle and nonmuscle cells from many different species of plants and
+animals. Unlike antibodies, the binding affinity does not change appreciably with
+actin from different species or sources. Nonspecific staining is negligible, and the
+contrast between stained and unstained areas is extremely large. It has been reported
+that phallotoxins are unable to bind to monomeric G-actin.
+
+Phallotoxins shift the monomer/polymer equilibrium toward the polymer, lowering the
+critical concentration for polymerization up to 30-fold. Phallotoxins also stabilize
+F-actin, inhibiting depolymerization by cytochalasins, potassium iodide, and elevated
+temperatures.
+
+Because the phallotoxin conjugates are small, with an approximate diameter of 12–15 Å
+and molecular weight of <2000 daltons, a variety of actin-binding proteins—including
+myosin, tropomyosin, troponin, and DNase I—can still bind to actin after treatment
+with phallotoxins. Even more significantly, phallotoxin-labeled actin filaments
+remain functional; labeled glycerinated muscle fibers still contract, and labeled
+actin filaments still move on solid-phase myosin substrates.
+
+Fluorescent phallotoxins can also be used to quantitate the amount of F-actin in
+cells. The unlabeled phallotoxins may be used as controls in blocking F-actin
+staining or in promoting actin polymerization. Our biotin-XX phalloidin allows
+researchers to visualize actin filaments by electron microscopy using standard
+enzyme-mediated avidin/streptavidin techniques. Eosin phalloidin can potentially be
+used for correlated fluorescence and electron microscopic studies. Deerinck and
+co-workers reported that when eosin conjugates are excited in the presence of
+diaminobenzidine (DAB), an insoluble, electron-dense DAB oxidation product is formed
+that is easily visualized with either light or electron microscopy.
+
+Even though phallotoxins have an LD50 of approximately 2 mg/kg when injected into
+the mouse, the toxins of A. phalloides that are responsible for most of the symptoms
+and fatalities associated with poisoning by this mushroom are the structurally related
+amatoxins. The major in vivo lesions produced by injected phallotoxin occur in the
+liver and are associated with stabilization of polymeric actin. Phallotoxins, however,
+tend not to be absorbed by the gastrointestinal tract even though they are stable
+between pH 3 and pH 9.
+
+
+MATERIALS
+================================================================================
+
+Storage and Safety
+
+Once reconstituted in methanol, the stock solutions are stable for at least one year
+when stored frozen at –20°C, desiccated, and protected from light. It appears that
+NBD phallacidin, and possibly all of these toxins, exhibit a small loss of activity
+when stored in aqueous solution at 2–6°C for over three weeks.
+
+While the amount of toxin present in a vial could be lethal only to a mosquito (LD50
+of phalloidin = 2 mg/kg), it should be handled with care.
+
+Materials Required but Not Provided
+
+• Methanol
+• PBS
+• Methanol-free formaldehyde
+• Acetone or Triton X-100
+• Fluorescent or enzyme-conjugated streptavidin, as prepared in step 1.9, for use
+  with biotin-XX phalloidin only
+• BSA, optional
+• Image-iT FX Signal Enhancer (I36933), optional
+• lysopalmitoylphosphatidylcholine, optional
+
+Preparing the Stock Solution
+
+Fluorescent Phallotoxins
+
+The vial contents should be dissolved in 1.5 mL methanol to yield a final
+concentration of 200 units/mL, which is equivalent to approximately 6.6 µM. One unit
+of phallotoxin is defined as the amount of material used to stain one microscope
+slide of fixed cells, according to the following protocol (see step 1.6), and is
+equivalent to 5 µL of methanolic stock solution for the fluorescent phallotoxins.
+
+Biotin-XX Phalloidin
+
+Staining cells with biotin-XX phalloidin (B7474) requires 1) the use of a higher
+concentration of the phallotoxin conjugate than when staining with fluorescent
+phallotoxins and 2) the addition of a fluorescent or enzyme-conjugated avidin or
+streptavidin detection reagent (see step 1.9). The vial contents should be dissolved
+in 0.5 mL methanol to yield a final concentration of 100 units/mL, which is
+equivalent to approximately 20 µM. For biotin-XX phalloidin, 10 µL of the
+methanolic stock solution is equivalent to one unit of phallotoxin, which is defined
+as the amount of material used to stain one microscope slide of fixed cells
+according to the following protocol (see step 1.6).
+
+Unlabeled Phalloidin
+
+Solutions of this product should be prepared just like the fluorescent phallotoxins
+described in Fluorescent Phallotoxins, taking into account the larger quantity of
+material provided.
+
+
+PROTOCOL
+================================================================================
+
+The procedure below was originally developed for use with NBD phallacidin. It has
+been successfully used with all of Invitrogen Molecular Probes phallotoxin
+conjugates. This procedure may not be optimum for a particular experimental system,
+but has yielded consistent results in most instances. The following protocol
+describes the staining procedure for adherent cells grown on glass coverslips.
+
+1. Formaldehyde-Fixed Cells
+
+   1.1. Wash cells twice with prewarmed phosphate-buffered saline, pH 7.4 (PBS).
+
+   1.2. Fix the sample in 3.7% formaldehyde solution in PBS for 10 minutes at room
+        temperature.
+
+        Note: Methanol can disrupt actin during the fixation process. Therefore,
+        it is best to avoid any methanol containing fixatives. The preferred
+        fixative is methanol-free formaldehyde.
+
+   1.3. Wash two or more times with PBS.
+
+   1.4. Place each coverslip in a glass petri dish and extract it with a solution
+        of acetone at –20°C or 0.1% Triton X-100 in PBS for 3 to 5 minutes.
+
+   1.5. Wash two or more times with PBS.
+
+   1.6. When staining with any of the fluorescent phallotoxins, dilute 5 µL
+        methanolic stock solution into 200 µL PBS for each coverslip to be
+        stained. To reduce nonspecific background staining with these conjugates,
+        add 1% bovine serum albumin (BSA) to the staining solution. It may also be
+        useful to pre-incubate fixed cells with PBS containing 1% BSA or with the
+        Image-iT FX signal enhancer (I36933) for 20–30 minutes prior to adding
+        the phallotoxin staining solution. When staining with biotin-XX phalloidin
+        (B7474), dilute 10 µL of the methanolic stock solution into 200 µL PBS
+        for each coverslip to be stained. When staining more than one coverslip,
+        adjust volumes accordingly. For a stronger signal, use 2 or 3 units per
+        coverslip.
+
+   1.7. Place the staining solution on the coverslip for 20 minutes at room
+        temperature (generally, any temperature between 4°C and 37°C is suitable).
+        To avoid evaporation, keep the coverslips inside a covered container
+        during the incubation.
+
+   1.8. Wash two or more times with PBS.
+
+   1.9. When using biotin-XX phalloidin, incubate for 30 minutes with 100 µL of a
+        10 µg/mL solution of fluorescent or enzyme-conjugated streptavidin
+        dissolved in 100 mM Tris-HCl, pH 7.5, 150 mM NaCl, 0.3% Triton X-100 and
+        1% BSA. Incubate for 15 minutes at room temperature. After incubation,
+        wash the coverslip with PBS. To develop enzyme activity, follow a
+        procedure recommended for the specific enzyme.
+
+   1.10. For long-term storage, the cells should be air dried and then mounted in a
+         permanent mountant such as ProLong Gold reagent or Cytoseal. Specimens
+         prepared in this manner retain actin staining for at least six months
+         when stored in the dark at 2–6°C.
+
+2. Simultaneous Fixation, Permeabilization, and Fluorescent Phallotoxin Staining
+
+   The phallotoxins appear to be stable for short periods in 4% formaldehyde
+   fixation buffers. This permits a rapid one-step fixation, permeabilization,
+   and labeling procedure as follows.
+
+   2.1. Prepare a 1 mL solution containing 50 to 100 µg/mL
+        lysopalmitoylphosphatidylcholine and 3.7% formaldehyde and then add 5–10
+        units of fluorescent phallotoxin (approximately 25 to 50 µL of methanolic
+        stock solution).
+
+   2.2. Place this staining solution on cells and incubate for 20 minutes at 4°C.
+
+   2.3. Rapidly wash three times with buffer.
+
+   2.4. Mount coverslips and view.
+
+
+TIPS FOR FLUORESCENCE MICROSCOPY
+================================================================================
+
+Photostability or resistance to photobleaching is a primary concern when making
+fluorescence measurements. Invitrogen brand product lines such as Alexa Fluor,
+Oregon Green, BODIPY, and rhodamine fluorophores (including Texas Red™-X) are
+significantly more photostable than NBD and fluorescein and will therefore enable
+more accurate photographic measurements.
+
+To further reduce photobleaching, minimize the exposure of fluorescently labeled
+specimens to light with neutral density filters and expose samples only when
+observing or recording a signal. Maximize collection of fluorescence by using a
+minimum of optics, high–numerical aperture objectives, relatively low magnification,
+high-quality optical filters, and high-speed film or high-efficiency detectors.
+
+Invitrogen Antifade reagents, including Molecular Probes SlowFade Gold and ProLong
+Gold antifade reagents, can extend the useful lives of many fluorescent probes.
+These reagents are also available in a formulation with DAPI. They can be used on
+fixed cell preparations but are not compatible with living cells. Cytoseal also
+appears to protect BODIPY fluorophores from photobleaching.
+
+
+REFERENCES
+================================================================================
+
+1. Wieland, T. in Phallotoxins, Springer-Verlag, New York (1986).
+2. J Muscle Res Cell Motil 9, 370 (1988).
+3. Methods Enzymol 85, 514 (1982).
+4. Eur J Biochem 165, 125 (1987).
+5. J Cell Biol 105, 1473 (1987).
+6. Nature 326, 805 (1987).
+7. Proc Natl Acad Sci USA 83, 6272 (1986).
+8. Blood 69, 945 (1987).
+9. Anal Biochem 200, 199 (1992).
+10. J Cell Biol 126, 901 (1994).
+11. Proc Natl Acad Sci USA 77, 980 (1980).
+12. Nature 284, 405 (1980).
+13. CRC Crit Rev Biochem 5, 185 (1978).
+14. J Cell Biol 106, 1229 (1988).
+15. J Cell Biol 103, 265a (1986).
+16. Eur J Cell Biol 24, 176 (1981).
+17. Proc Natl Acad Sci USA 74, 5613 (1977).

biomni/tool/protocols/thermofisher/ActinGreen 488 ReadyProbes Reagent Protocol _ Thermo Fisher Scientific - US.txt

@@ -0,0 +1,67 @@
+PROTOCOL: ActinGreen 488 ReadyProbes Reagent Protocol
+SOURCE: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/protocols/actin-green-488-readyprobes-protocol.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+F-actin stain for fixed cells
+
+ActinGreen 488 ReadyProbes Reagent is a selective, high-affinity F-actin probe conjugated to our bright, photostable green-fluorescent Alexa Fluor 488 dye. Like other ReadyProbes reagents, the ActinGreen 488 reagent is formulated as a room temperature–stable solution and comes in a convenient dropper bottle.
+
+APPLICATIONS
+================================================================================
+
+This protocol can be used for:
+• F-actin staining in fluorescence microscopy
+
+This protocol should not be used for:
+• Flow cytometry
+
+MATERIALS REQUIRED
+================================================================================
+
+• Cells growing in culture
+• ActinGreen 488 ReadyProbes Reagent (Cat. No. R37110)
+• Fluorescence microscope
+
+PROTOCOL
+================================================================================
+
+Labeling fixed cells
+
+First, fix and permeabilize cultured cells with a protocol appropriate for your sample. Counterstain as desired. DAPI may be added directly to fixative.
+
+1. Wash the cells 1–3 times in PBS as needed.
+2. Add two drops of ActinGreen 488 ReadyProbes Reagent per milliliter of medium.
+3. Incubate for 30 minutes, protected from light.
+4. Remove the stain solution.
+5. Wash the cells 2–3 times in PBS.
+6. Image the cells.
+
+SPECTRAL INFORMATION AND STORAGE
+================================================================================
+
+ActinGreen 488:
+• Excitation/Emission (nm): 495/518
+• Standard filter set: FITC
+• EVOS Light Cube: GFP
+• Storage conditions: Room temperature
+
+PROTOCOL TIPS
+================================================================================
+
+• In some cases, more or fewer drops may be needed to achieve optimal staining intensity.
+• Image quality may be improved by replacing the culture medium with Live Cell Imaging Solution (Cat. No. A14291DJ).
+
+EXAMPLE APPLICATION
+================================================================================
+
+Multicolor staining of HeLa cells.
+
+HeLa cells were grown on coverslips and were fixed and permeabilized using the Image-iT Fixation/Permeabilization Kit. Cells were then stained to detect actin (green fluorescence from ActinGreen 488 ReadyProbes Reagent), B-catenin (red-orange fluorescence), and nuclei (blue fluorescence).
+
+PRODUCT CATALOG NUMBER
+================================================================================
+
+R37110

biomni/tool/protocols/thermofisher/ActinRed 555 ReadyProbes Reagent Protocol _ Thermo Fisher Scientific - US.txt

@@ -0,0 +1,67 @@
+PROTOCOL: ActinRed 555 ReadyProbes Reagent Protocol
+SOURCE: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/protocols/actin-red-555-readyprobes-protocol.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+F-actin stain for fixed cells
+
+ActinRed 555 ReadyProbes Reagent is a selective, high-affinity F-actin probe conjugated to the red-orange fluorescent dye tetramethylrhodamine (TRITC). Like other ReadyProbes reagents, ActinRed 555 reagent is formulated as a room temperature–stable solution and comes in a convenient dropper bottle.
+
+APPLICATIONS
+================================================================================
+
+This protocol can be used for:
+• F-actin staining in fluorescence microscopy
+
+This protocol should not be used for:
+• Flow cytometry
+
+MATERIALS REQUIRED
+================================================================================
+
+• Cells growing in culture
+• ActinRed 555 ReadyProbes Reagent (Cat. No. R37112)
+• Fluorescence microscope
+
+PROTOCOL
+================================================================================
+
+Labeling fixed cells
+
+First, fix and permeabilize cultured cells with a protocol appropriate for your sample. Counterstain as desired. DAPI may be added directly to fixative.
+
+1. Wash the cells 1–3 times in PBS as needed.
+2. Add two drops of ActinRed 555 ReadyProbes Reagent per milliliter of medium.
+3. Incubate for 30 minutes, protected from light.
+4. Remove the stain solution.
+5. Wash the cells 2–3 times in PBS.
+6. Image the cells.
+
+SPECTRAL INFORMATION AND STORAGE
+================================================================================
+
+ActinRed 555:
+• Excitation/Emission (nm): 540/565
+• Standard filter set: TRITC
+• EVOS Light Cube: TRITC
+• Storage conditions: Room temperature
+
+PROTOCOL TIPS
+================================================================================
+
+• In some cases, more or fewer drops may be needed to achieve optimal staining intensity.
+• Image quality may be improved by replacing the culture medium with Live Cell Imaging Solution (Cat. No. A14291DJ).
+
+EXAMPLE APPLICATION
+================================================================================
+
+Multicolor staining of BPAE cells.
+
+BPAE cells were fixed and permeabilized using the Image-iT Fixation/Permeabilization Kit. Actin was stained using ActinRed 555 ReadyProbes Reagent, and nuclei were stained with NucBlue Fixed Cell ReadyProbes Reagent.
+
+PRODUCT CATALOG NUMBER
+================================================================================
+
+R37112

biomni/tool/protocols/thermofisher/Adherent Cell Culture Protocol _ Thermo Fisher Scientific - US.txt

@@ -0,0 +1,75 @@
+PROTOCOL: Adherent Cell Culture Protocol
+SOURCE: https://www.thermofisher.com/us/en/home/references/gibco-cell-culture-basics/cell-culture-protocols/subculturing-adherent-cells.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+An adherent cell culture is characterized by the cell's ability to proliferate when attached to a growth-promoting substrate, a property known as "anchorage dependence." The following protocols detail various methods for dissociation of adherent cells from cultureware and subsequent procedures for subculturing, or splitting, adherent cells.
+
+MATERIALS FOR DETACHING AND PASSAGING ADHERENT CELLS
+================================================================================
+
+In addition to the catalog items below, detaching and passaging adherent cells also requires the following:
+
+• Adherent cells
+• Tissue-culture treated flasks, plates or dishes
+• Complete growth medium: basal medium, serum, and supplements pre-warmed to 37°C
+• Disposable, sterile 15 mL tubes
+• 37°C incubator with humidified atmosphere of 5% CO₂
+• Pipettes and pipette tips
+• Automated cell counter such as the Countess Automated Cell Counter
+
+Catalog numbers: 13151014, 13150016, 14190144, 15400054, 15090046, 12604013, 15575020, 179693, 17105041, 15040066, 15250061
+
+PROTOCOL FOR PASSAGING ADHERENT CELLS
+================================================================================
+
+The following adherent cell culture protocol describes how to subculture mammalian cells. Note that the procedure for subculturing insect cells differs from that for mammalian cells in several crucial steps. For more information, refer to "Considerations for passaging adherent insect cells" below.
+
+For passaging a particular cell line within your experiments, we recommend that you closely follow the instructions provided with each product you use, especially regarding formulation of your complete growth medium. The consequences of deviating from the culture conditions specified for a particular cell type can range from the expression of aberrant phenotypes to a complete failure of the cell culture. The SDSs for all Thermo Fisher Scientific products can be accessed here.
+
+Before passaging, ensure the solutions and equipment that come in contact with the cells are sterile. Use proper sterile technique throughout the subculture procedure and work in a laminar flow hood.
+
+Routinely monitor your cell's viability prior to subculturing. Adherent cells should be passaged at log phase with viability greater than 90% at the time of subculturing.
+
+1. Remove and discard the spent cell culture media from the culture vessel.
+
+2. Wash cells using a balanced salt solution without calcium and magnesium (approximately 2 mL per 10 cm² culture surface area). Gently add wash solution to the side of the vessel opposite the attached cell layer to avoid disturbing the cell layer and rock the vessel back and forth several times. The wash step removes any traces of serum, calcium, and magnesium that would inhibit the action of the dissociation reagent.
+
+3. Remove and discard the wash solution from the culture vessel.
+
+4. Add the pre-warmed dissociation reagent such as trypsin or TrypLE to the side of the flask; use enough reagent to cover the cell layer (approximately 0.5 mL per 10 cm²). Gently rock the container to get complete coverage of the cell layer.
+
+5. Incubate the culture vessel at room temperature for approximately 2 minutes. Note that the actual incubation time varies with the cell line used.
+
+6. Observe the cells under the microscope for detachment. If cells are less than 90% detached, increase the incubation time a few more minutes, checking for dissociation every 30 seconds. You may also tap the vessel to expedite cell detachment.
+
+7. When ≥ 90% of the cells have detached, tilt the vessel for a minimal length of time to allow the cells to drain. Add the equivalent of 2 volumes (twice the volume used for the dissociation reagent) of pre-warmed complete growth medium. Disperse the medium by pipetting over the cell layer surface several times.
+
+8. Transfer the cells to a 15 mL conical tube and centrifuge them at 200 x g for 5 to 10 minutes. Note that the centrifuge speed and time vary based on the cell type.
+
+9. Resuspend the cell pellet in a minimal volume of pre-warmed complete growth medium and remove a sample for counting.
+
+10. Determine the total number of cells and percent viability using a hemocytometer and the Trypan blue exclusion protocol, or the Invitrogen Countess Automated Cell Counter. If necessary, add growth media to the cells to achieve the desired cell concentration and recount the cells.
+
+11. Dilute cell suspension to the seeding density recommended for the cell line, and pipet the appropriate volume into new cell culture vessels and return the cells to the incubator. If using culture flasks, loosen the caps before placing them in the incubator to allow proper gas exchange unless you are using vented flasks with gas-permeable caps.
+
+CONSIDERATIONS FOR PASSAGING ADHERENT INSECT CELLS
+================================================================================
+
+While the general procedure for subculturing insect cells follows the same steps as mammalian cells, some key requirements of these culture systems are different. For the best results, always follow the instructions provided with each product you are using in your experiments.
+
+When to subculture adherent insect cells
+
+Passage insect cells at log phase, before they reach confluency, unless your cells are strongly adherent. If your insect cells are strongly adherent, you may passage them at confluency or slightly after when they are starting to pull away from the bottom of the flask because they will be easier to dislodge. However, insect cells that are repeatedly passaged at densities past confluency display decreased doubling times, decreased viability, and a decreased ability to attach.
+
+In contrast, passaging insect cells in adherent culture before they reach confluency requires more mechanical force to dislodge them from the monolayer. When repeatedly subcultured before confluency, these cells also display decreased doubling times, decreased viability, and are considered unhealthy. Densities lower than 20% confluency inhibit cell growth as well.
+
+How to subculture insect cells
+
+• Maintain insect cells at 27°C in a non-humidified environment. Cells can be maintained at room temperature on the bench top if protected from light or in a drawer. However, a 27°C controlled environment is recommended.
+• CO₂ exchange is not recommended for insect cell culture.
+• Use media specifically formulated for insect cell growth. Insect cells are cultured in growth media that are usually more acidic than those used for mammalian cells such as Grace's medium.
+• Insect cells attach very tightly to substrates under serum-free conditions and require additional effort to detach. To dislodge the cells, you may need to give the flask one quick shake using a wrist-snapping motion. To avoid contamination, always tighten the cap before this procedure.
+• Do not shake the flask vigorously, because it may result in damage to the cells.

biomni/tool/protocols/thermofisher/Basic Sandwich ELISA Protocol _ Thermo Fisher Scientific - US.txt

@@ -0,0 +1,98 @@
+PROTOCOL: Basic Sandwich ELISA Protocol
+SOURCE: Thermo Fisher Scientific - US
+URL: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/elisa-protocol/general-elisa-protocol.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+The sandwich ELISA (enzyme-linked immunosorbent assay) is a highly sensitive immunoassay technique used to detect and quantify specific proteins or antigens in samples. This method uses two antibodies: a capture antibody that binds to the target antigen, and a detection antibody that recognizes a different epitope on the same antigen. The detection antibody is conjugated to an enzyme (such as horseradish peroxidase or alkaline phosphatase) that produces a measurable signal.
+
+MATERIALS REQUIRED
+================================================================================
+
+• 96-well microplate (pre-coated with capture antibody or uncoated for manual coating)
+• Capture antibody (specific to target antigen)
+• Detection antibody (conjugated to HRP or AP, specific to target antigen)
+• Blocking buffer (e.g., 1% BSA or 5% non-fat dry milk in PBS)
+• Wash buffer (PBS with 0.05% Tween-20, PBST)
+• Sample diluent (PBS or assay buffer)
+• Standard/reference antigen (for standard curve)
+• Substrate solution (TMB for HRP, or pNPP for AP)
+• Stop solution (2N H2SO4 for TMB, or 3N NaOH for pNPP)
+• Plate reader (capable of reading absorbance at appropriate wavelength)
+
+PROTOCOL
+================================================================================
+
+Day 1: Plate Coating (if using uncoated plates)
+
+1. Dilute capture antibody in coating buffer (typically 0.1 M carbonate-bicarbonate buffer, pH 9.6, or PBS) to the recommended concentration (usually 1-10 µg/mL).
+
+2. Add 100 µL of diluted capture antibody to each well of a 96-well microplate.
+
+3. Cover plate and incubate overnight at 4°C, or for 2 hours at room temperature.
+
+4. Remove the coating solution and wash plate 3 times with wash buffer (200-300 µL per well). Invert plate and tap on paper towels to remove excess liquid.
+
+Day 2: Blocking and Assay
+
+5. Add 200-300 µL of blocking buffer to each well.
+
+6. Incubate for 1-2 hours at room temperature, or overnight at 4°C.
+
+7. Wash plate 3 times with wash buffer as described in step 4.
+
+8. Prepare samples and standards:
+   - Dilute samples in sample diluent to appropriate concentration
+   - Prepare standard curve by serial dilution of reference antigen in sample diluent
+   - Include blank wells with sample diluent only
+
+9. Add 100 µL of samples and standards to appropriate wells in duplicate or triplicate.
+
+10. Cover plate and incubate for 1-2 hours at room temperature, or overnight at 4°C.
+
+11. Wash plate 3-5 times with wash buffer as described in step 4.
+
+12. Dilute detection antibody in blocking buffer or antibody diluent to recommended concentration (typically 0.5-2 µg/mL).
+
+13. Add 100 µL of diluted detection antibody to each well.
+
+14. Cover plate and incubate for 1-2 hours at room temperature.
+
+15. Wash plate 3-5 times with wash buffer as described in step 4.
+
+16. Prepare substrate solution according to manufacturer's instructions.
+
+17. Add 100 µL of substrate solution to each well.
+
+18. Incubate at room temperature, protected from light, for 5-30 minutes (or until desired color development).
+
+19. Add 50-100 µL of stop solution to each well (if using TMB or other substrates requiring stopping).
+
+20. Read plate immediately using a microplate reader at the appropriate wavelength:
+    - TMB: 450 nm (with 570 nm or 630 nm reference)
+    - pNPP: 405 nm
+    - OPD: 492 nm
+
+DATA ANALYSIS
+================================================================================
+
+1. Subtract the average absorbance of blank wells from all sample and standard values.
+
+2. Plot standard curve: absorbance (y-axis) vs. concentration (x-axis) using standard values.
+
+3. Use the standard curve to determine the concentration of unknown samples.
+
+4. For accurate results, ensure sample values fall within the linear range of the standard curve.
+
+PROTOCOL TIPS
+================================================================================
+
+• Always include negative controls (no antigen) and positive controls (known concentration of antigen)
+• Use fresh substrate solutions and prepare just before use
+• Avoid bubbles when adding solutions to wells
+• Ensure consistent incubation times and temperatures
+• Wash thoroughly between steps to reduce background signal
+• Optimize antibody concentrations and incubation times for your specific assay
+• For best results, use high-quality, validated antibody pairs

biomni/tool/protocols/thermofisher/BestProtocols_ Annexin V Staining Protocol for Flow Cytometry _ Thermo Fisher Scientific - US.txt

@@ -0,0 +1,128 @@
+PROTOCOL: BestProtocols: Annexin V Staining Protocol for Flow Cytometry
+SOURCE: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/protocols/annexin-v-staining-flow-cytometry.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+Note: This protocol is intended for use with the specific products mentioned within it. Substituting different products is not recommended.
+
+The use of the annexin V apoptosis assay protocol is a common method for detecting apoptotic cells. The below protocols are recommended for use with the specific flow cytometry kits mentioned. Please see the Annexin V Staining page for a discussion about general experimental conditions and avoiding false positives, or to review a selection guide for all of our annexin V products.
+
+GENERAL NOTES
+================================================================================
+
+• Invitrogen Fixable Viability Dye (FVD) eFluor 450 is not recommended for use with Annexin V Apoptosis Detection Kits.
+• Due to the calcium dependence of the Annexin V:PS interaction, it is critical to avoid buffers containing EDTA or other calcium chelators during Annexin V experiments.
+• Annexin V can only be used as a marker of apoptosis in cells where the plasma membrane is intact. Destroying the integrity of the plasma membrane will allow binding of Annexin V to PS inside the cell.
+
+PROTOCOL A: ANNEXIN V STAINING PROTOCOL
+================================================================================
+
+Materials
+
+• 12 x 75 mm round-bottom tubes
+• 1X PBS
+• Annexin V Apoptosis Detection kit (any one of the kits listed). Each kit includes an Annexin V conjugate:
+  - eFluor 450 (Cat. Nos. 88-8006-72, 88-8006-74)
+  - FITC (Cat. Nos. 88-8005-72, 88-8005-74)
+  - PerCP-eFluor 710 (Cat. Nos. 88-8008-72, 88-8008-74)
+  - PE (Cat. Nos. 88-8102-72, 88-8102-74)
+  - PE-Cyanine7 (Cat. Nos. 88-8103-72, 88-8103-74)
+  - APC (Cat. Nos. 88-8007-72, 88-8007-74)
+• 10X Binding buffer
+
+Note: PerCP-eFluor 710 (Cat. Nos. 88-8008-72, 88-8008-74) do not include a viability dye. We recommend using this format in combination with a Invitrogen Fixable Viability Dye (FVD) such as FVD eFluor 660 (Cat. Nos. 65-0864-14, 65-0864-18), FVD eFluor 506 (Cat. Nos. 65-0866-14, 65-0866-18), or FVD eFluor 780 (Cat. Nos. 65-0865-14, 65-8065-18).
+
+Experimental procedure
+
+1. Prepare 1X binding buffer by mixing 1 part of 10X binding buffer with 9 parts of distilled water.
+2. Harvest cells.
+3. Wash cells once in 1X PBS, then once in 1X binding buffer.
+4. Resuspend cells in 1X Binding Buffer at 1-5 x 10⁶ cells/mL.
+5. Add 5 μL of fluorochrome-conjugated Annexin V to 100 μL of the cell suspension.
+6. Incubate 10-15 minutes at room temperature. Protect from light.
+7. Add 2 mL 1X binding buffer and centrifuge at 400-600 x g for 5 minutes at room temperature. Discard supernatant.
+8. Resuspend cells in 200 µL of 1X binding buffer.
+9. Add 5 μL of Propidium Iodide Staining Solution or 7-AAD Viability Staining Solution and incubate 5-15 minutes on ice or at room temperature.
+
+Note: Propidium iodide and 7-AAD must remain in the buffer during acquisition. Do not wash cells after the addition of propidium iodide or 7-AAD.
+
+10. Analyze by flow cytometry.
+
+Note: Cells should be analyzed within 4 hours after the initial incubation period due to adverse effects on the viability of cells left in the presence of propidium iodide or 7-AAD for prolonged periods. Store at 2–8°C and protect from light until ready for analysis.
+
+PROTOCOL B: ANNEXIN V STAINING PROTOCOL WITH FIXABLE VIABILITY DYES
+================================================================================
+
+Materials
+
+• 12 x 75 mm round-bottom tubes
+• 1X PBS
+• Annexin V Apoptosis Detection kit (any one of the kits listed). Each kit includes an Annexin V conjugate:
+  - eFluor 450 (Cat. Nos. 88-8006-72, 88-8006-74)
+  - FITC (Cat. Nos. 88-8005-72, 88-8005-74)
+  - PerCP-eFluor 710 (Cat. Nos. 88-8008-72, 88-8008-74)
+  - PE (Cat. Nos. 88-8102-72, 88-8102-74)
+  - PE-Cyanine7 (Cat. Nos. 88-8103-72, 88-8103-74)
+  - APC (Cat. Nos. 88-8007-72, 88-8007-74)
+• 10X Binding buffer
+• Flow Cytometry Staining Buffer (Cat. No. 00-4222)
+• PBS (azide- and serum/protein-free PBS)
+• FVD eFluor 660 (Cat. Nos. 65-0864-14, 65-0864-18), FVD eFluor 506 (Cat. Nos. 65-0866-14, 65-0866-18) or FVD eFluor 780 (Cat. Nos. 65-0865-14, 65-8065-18)
+
+Note: FVD eFluor 450 is not recommended for use with the Annexin V Apoptosis Detection Kits.
+
+Experimental procedure
+
+1. Prepare 1X binding buffer by mixing 1 part of 10X binding buffer with 9 parts of distilled water.
+2. Wash cells twice in azide-free and serum/protein-free PBS.
+3. Resuspend cells at 1-10 x 10⁶ cells/mL in azide-free and serum/protein-free PBS.
+4. Add 1 µL of FVD per 1 mL of cells and vortex immediately.
+5. Incubate for 30 minutes at 2-8°C. Protect from light.
+6. Wash cells twice with Flow Cytometry Staining Buffer or equivalent.
+7. Wash cells once with 1X Binding Buffer.
+8. Resuspend cells in 1X Binding Buffer at 1-5 x 10⁶ cells/mL.
+9. Add 5 μL of fluorochrome-conjugated Annexin V to 100 μL of the cell suspension.
+10. Incubate 10-15 minutes at room temperature. Protect from light.
+11. Add 2 mL of 1X binding buffer and centrifuge at 400-600 x g for 5 minutes at room temperature. Discard supernatant.
+12. Resuspend cells in 200 μL of 1X Binding Buffer.
+13. Analyze by flow cytometry.
+
+PROTOCOL C: ANNEXIN V STAINING PROTOCOL WITH SURFACE AND INTRACELLULAR STAINING
+================================================================================
+
+Materials
+
+• 12 x 75 mm round-bottom tubes
+• 1X PBS (azide- and serum/protein-free PBS)
+• Flow Cytometry Staining Buffer Set (Cat. No. 00-4222)
+• Foxp3/Transcription Factor Staining Buffer Set (Cat. No. 00-5523) or Intracellular Fixation & Permeabilization Buffer Set (Cat. No. 88-8824)
+• Annexin V Apoptosis Detection kit (any one of the kits listed). Each kit includes an Annexin V conjugate:
+  - eFluor 450 (Cat. Nos. 88-8006-72, 88-8006-74)
+  - FITC (Cat. Nos. 88-8005-72, 88-8005-74)
+  - PerCP-eFluor 710 (Cat. Nos. 88-8008-72, 88-8008-74)
+  - PE (Cat. Nos. 88-8102-72, 88-8102-74)
+  - PE-Cyanine7 (Cat. Nos. 88-8103-72, 88-8103-74)
+  - APC (Cat. Nos. 88-8007-72, 88-8007-74)
+• 10X Binding buffer
+• FVD eFluor 660 (Cat. Nos. 65-0864-14, 65-0864-18), FVD eFluor 506 (Cat. Nos. 65-0866-14, 65-0866-18) or FVD eFluor 780 (Cat. Nos. 65-0865-14, 65-8065-18)
+
+Note: FVD eFluor 450 is not recommended for use with the Annexin V Apoptosis Detection Kits.
+
+Experimental procedure
+
+1. Prepare 1X binding buffer by mixing 1 part of 10X binding buffer with 9 parts of distilled water.
+2. Stain cell surface antigen(s). Refer to 'Staining Cell Surface Targets, Protocol A'.
+3. Wash cells twice in azide-free and serum/protein-free PBS.
+4. Resuspend cells at 1-10 x 10⁶ cells/mL in azide-free and serum/protein-free PBS.
+5. Add 1 µL of Fixable Viability Dye per 1 mL of cells and vortex immediately.
+6. Incubate for 30 minutes at 2-8°C. Protect from light.
+7. Wash cells twice in Flow Cytometry Staining Buffer or equivalent.
+8. Wash cells once with 1X binding buffer.
+9. Resuspend cells in 1X binding buffer at 1-5 x 10⁶ cells/mL.
+10. Add 5 μL of fluorochrome-conjugated Annexin V to 100 μL of the cell suspension.
+11. Incubate 10-15 minutes at room temperature. Protect from light.
+12. Wash cells once with 1X binding buffer.
+13. Stain intracellular antigen(s). Refer to 'Staining Intracellular Antigens, Protocol A or B'.
+14. Analyze by flow cytometry.

biomni/tool/protocols/thermofisher/BestProtocols_ Cell Preparation for Flow Cytometry Protocols _ Thermo Fisher Scientific - US.txt

@@ -0,0 +1,135 @@
+PROTOCOL: BestProtocols: Cell Preparation for Flow Cytometry Protocols
+SOURCE: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/protocols/cell-preparation-flow-cytometry.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+Note: This protocol is intended for use with the specific products mentioned within it. Substituting different products is not recommended.
+
+Introduction
+
+Single-cell suspensions are required for all flow cytometry assays. Thus, peripheral blood cells or cells that grow in suspension are well suited for analysis by flow cytometry. Adherent cell lines, solid tissue samples, and tumors require processing into single-cell suspensions before they can be analyzed. Numerous protocols are available and may involve enzymatic digestion or mechanical dissociation of the tissue. Care should be used when chelation or enzymatic digestion are used, as these may result in the destruction of the antibody epitope. In all situations, removing cell clumps, dead cells, and debris is essential to eliminate false positives and obtain results of the highest quality.
+
+PROTOCOL A: TISSUE CULTURE CELLS
+================================================================================
+
+Materials
+
+• Invitrogen Accutase Enzyme Cell Detachment Medium (cat. no. 00-4555) or trypsin or ethylenediaminetetraacetic acid (EDTA)
+• Flow Cytometry Staining Buffer (cat. no. 00-4222)
+• 15- or 50-mL conical centrifuge tubes
+
+Experimental procedure
+
+1. For cells that grow in suspension, decant the cells into a conical centrifuge tube and perform a cell count and viability analysis. Proceed to Step 4.
+
+2. For adherent cells lines, detach cells from the plate using one of the following options:
+   - Cell scraper
+   - Trypsin
+   - EDTA (10 mM in phosphate-buffered saline)
+   - Accutase Enzyme Cell Detachment Medium
+
+3. Place cells into a conical centrifuge tube and use a Invitrogen Pipetman to dissociate any clumps, and then perform cell count and viability analysis.
+
+4. Centrifuge cells and resuspend in an appropriate volume of Flow Cytometry Staining Buffer so that the final cell concentration is 1 x 10⁷ cells/mL (other cell concentrations may be appropriate for different experiments).
+
+PROTOCOL B: LYMPHOID TISSUE
+================================================================================
+
+Mechanical disruption of lymphoid tissue is generally sufficient to release cells to a single cell suspension.
+
+Materials
+
+• 60 x 15 mm tissue culture dish
+• 3-mL syringe or frosted glass microscope slides
+• Cell strainer (nylon mesh)
+• Flow Cytometry Staining Buffer (cat. no. 00-4222) or other buffer of choice
+• 15- or 50-mL conical centrifuge tubes
+
+Experimental procedure
+
+Note: If cells are to be cultured, perform all steps using aseptic technique and buffers that do not contain azide.
+
+1. Harvest tissue (spleen; thymus; lymph nodes) into a tissue culture dish containing 10 mL of Flow Cytometry Staining Buffer or buffer of choice. Tease apart into a single-cell suspension by pressing with the plunger of a 3-mL syringe. Alternatively, mash tissue between the frosted ends of two microscope slides using 10 mL of Flow Cytometry Staining Buffer.
+
+2. Place a cell strainer on top of a 15- or 15-mL conical tube. Pass cells from the tissue culture dish through the cell strainer to eliminate clumps and debris.
+
+3. Centrifuge cell suspension at 300-400 x g for 4-5 minutes at 2-8°C. Discard the supernatant.
+
+4. Resuspend the cell pellet in an appropriate volume of Flow Cytometry Staining Buffer or buffer of choice and perform a cell count and viability analysis.
+
+5. Centrifuge cells as in Step 3 and resuspend in appropriate volume of Flow Cytometry Staining Buffer or buffer of choice so that the final cell concentration is 1 x 10⁷ cells/mL (other cell concentrations may be appropriate for different experiments).
+
+PROTOCOL C: NON-LYMPHOID TISSUE
+================================================================================
+
+Materials
+
+• Scissors or scalpel blade
+• Phosphate-buffered saline (PBS) or other suitable physiologic buffer
+• Enzymes for tissue digestion
+• 60 x 15 mm tissue culture dish
+• 3-mL syringe
+• Cell strainer (nylon mesh)
+• Flow Cytometry Staining Buffer (cat. no. 00-4222) or other buffer of choice
+• 15- or 50-mL conical centrifuge tubes
+
+Experimental procedure
+
+Note: If cells are to be cultured, perform all steps using aseptic technique and buffers that do not contain azide.
+
+1. Harvest tissue and mince into 2-4 mm pieces using scissors or scalpel blade.
+
+2. Add appropriate amount of enzyme(s) diluted in PBS and incubate at the optimal temperature for the appropriate amount of time according to enzyme manufacturer instructions.
+
+3. Disperse cells by gentle pipetting and filter through a cell strainer to eliminate clumps and debris. Collect cell suspension in a conical tube.
+
+4. Centrifuge cells at 300-400 x g for 4-5 minutes at 2-8°C. Discard the supernatant.
+
+5. Resuspend the cell pellet in PBS.
+
+6. Centrifuge cells as in Step 4.
+
+7. Repeat Steps 5 and 6.
+
+8. Resuspend the cell pellet in an appropriate volume of Flow Cytometry Staining Buffer or buffer of choice and perform a cell count and viability analysis.
+
+9. Centrifuge cells as in Step 4 and resuspend in appropriate volume of Flow Cytometry Staining Buffer or buffer of choice so that the final cell concentration is 1 x 10⁷ cells/mL (other cell concentrations may be appropriate for different experiments).
+
+PROTOCOL D: ISOLATION OF PBMC FROM WHOLE BLOOD
+================================================================================
+
+Materials
+
+• PBS
+• Invitrogen Ficoll Paque or other density separation medium
+• Flow Cytometry Staining Buffer (cat. no. 00-4222) or other buffer of choice
+• 15- or 50-mL conical centrifuge tubes
+
+Experimental procedure
+
+Note: If cells are to be cultured, perform all steps using aseptic technique and buffers that do not contain azide.
+
+1. Dilute blood sample at least 1:1 with PBS in a conical tube.
+
+2. Underlay the diluted sample with a volume of Ficoll that is equal to the original sample volume.
+
+3. Centrifuge at 400 x g for 20 minutes at room temperature with the brake OFF.
+
+4. Harvest PBMC located at the interface of the PBS and Ficoll layers into a fresh tube.
+
+5. Fill the tube with PBS to wash the cells.
+
+6. Centrifuge the cells at 300-400 x g for 4-5 minutes at 2-8°C. Discard supernatant.
+
+7. Resuspend the cell pellet in an appropriate volume of Flow Cytometry Staining Buffer or buffer of choice and perform a cell count and viability analysis.
+
+8. Centrifuge cells as in Step 4 and resuspend in appropriate volume of Flow Cytometry Staining Buffer or buffer of choice so that the final cell concentration is 1 x 10⁷ cells/mL (other cell concentrations may be appropriate for different experiments).
+
+USEFUL RESOURCES
+================================================================================
+
+Worthington tissue dissociation guide (https://www.worthington-biochem.com/tools-resources/tissue-dissociation-guide)
+
+The Worthington Tissue Dissociation Guide provides a useful summary and guide of the various methods that can be used for tissue dissociation.

biomni/tool/protocols/thermofisher/BrdU Labeling & Detection Protocol _ Thermo Fisher Scientific - US.txt

@@ -0,0 +1,93 @@
+PROTOCOL: BrdU Labeling & Detection Protocol
+SOURCE: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/protocols/brdu-labeling-and-detection-protocol.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+BrdU (5-bromo-2'-deoxyuridine) is a thymidine analog used in DNA synthesis–based cell proliferation assays. The BrdU assay principle involves the incorporation of BrdU into newly synthesized DNA during cell division. BrdU labeling and detection are achieved by using an anti-BrdU antibody, which binds to the incorporated BrdU, allowing for the imaging and quantification of proliferating cells. This process, often referred to as BrdU staining/labeling, is a critical component of the BrdU cell proliferation assay protocol and is detailed here.
+
+APPLICATIONS
+================================================================================
+
+This protocol can be used for:
+• Detecting DNA synthesis using a fluorescence microscope
+
+This protocol should not be used for:
+• Flow cytometry (see BrdU Staining Kit Protocol for Flow Cytometry)
+
+MATERIALS REQUIRED
+================================================================================
+
+You will need the following for this protocol:
+
+• Proliferating adherent cells in an appropriate vessel for imaging
+• BrdU (5-Bromo-2´-Deoxyuridine)
+• Invitrogen Anti-BrdU monoclonal antibody (find additional anti-brdu antibody options)
+• Invitrogen Alexa Fluor–labeled secondary antibody, such as Goat anti-Mouse IgG (H+L) Cross-Adsorbed Secondary Antibody, Alexa Fluor 488
+• Complete tissue culture medium, such as DMEM
+• DPBS or similar wash solution
+• Fixative (i.e., 3.7% formaldehyde in PBS)
+• Permeabilization buffer, such as 0.1% Invitrogen Triton X-100 in PBS
+• 12 N HCl
+• Phosphate/citric acid buffer, pH 7.4 (182 mL of 0.2 M Na₂HPO₄ + 18 mL 0.1 M citric acid)
+• Antibody stain solution (PBS/0.1% Triton/5% normal goat serum)
+• Ice bucket
+• Fluorescence microscope
+
+PROTOCOL
+================================================================================
+
+Prepare stock solutions
+
+1. Dissolve 100 mg BrdU in 32.5 mL anhydrous DMSO (10 mM stock solution).
+2. Dilute 10 µL of this stock solution in 10 mL of 37°C tissue culture medium to make a 10 µM labeling solution.
+
+Label cells with BrdU
+
+1. Culture cells in appropriate vessel for microscopy.
+2. Remove culture medium from cells and replace with BrdU labeling solution.
+3. Incubate cells at 37°C for 2 hours.
+4. Remove labeling solution and wash two times with PBS.
+5. Wash with PBS (3 times, 2 minutes each).
+
+Fix, permeabilize, and acid-wash
+
+1. Remove PBS and add 1 mL of 3.7% formaldehyde in PBS to each well.
+2. Incubate for 15 minutes at room temperature.
+3. Wash with PBS (3 times, 2 minutes each).
+4. Remove PBS and add 1 mL of Triton X-100 permeabilization buffer to each well.
+5. Incubate for 20 minutes at room temperature.
+6. Remove permeabilization buffer and add 1 mL of 1 N HCl.
+7. Incubate 10 minutes on ice.
+8. Remove this solution and add 1 mL of 2 N HCl.
+9. Incubate 10 minutes at room temperature.
+10. Add 1 mL phosphate/citric acid buffer, pH 7.4.
+11. Incubate 10 minutes at room temperature.
+12. Wash with Triton X-100 permeabilization buffer (3 times, 2 minutes each).
+
+Detect incorporated BrdU
+
+1. Remove this solution and add 1 mL of antibody staining buffer.
+2. Add anti-BrdU primary antibody.
+3. Incubate overnight at room temperature.
+4. Wash with Triton X-100 permeabilization buffer (3 times, 2 minutes each).
+5. Add fluorescently labeled secondary antibody.
+6. Incubate one hour at room temperature.
+
+Imaging
+
+1. Add PBS to each well.
+2. Image cells with appropriate filters.
+
+PROTOCOL TIPS
+================================================================================
+
+• Leftover BrdU stock solution can be stored frozen for up to one year.
+• Use short incubation time for rapidly proliferating cells, and longer incubation for slow-growing cells.
+• Acid treatment separates DNA into single strands so the primary antibody can access the incorporated BrdU.
+
+PRODUCT CATALOG NUMBERS
+================================================================================
+
+B23151, B35130, A11001, 14040141, HFH10

biomni/tool/protocols/thermofisher/BrdU Staining Protocol for Flow Cytometry _ Thermo Fisher Scientific - US.txt

@@ -0,0 +1,85 @@
+PROTOCOL: BrdU Staining Protocol for Flow Cytometry
+SOURCE: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/protocols/brdu-staining-flow-cytometry.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+The BrdU Staining Kits for flow cytometry provide all necessary reagents and buffers for identifying and analyzing proliferating cells using flow cytometry. BrdU, or 5-bromo-2'-deoxyuridine, is a synthetic analog of thymidine that integrates into newly synthesized DNA during the S-phase of mitosis. This kit allows for the staining of BrdU-incorporated DNA, facilitating the examination of cell proliferation alongside other cell surface or intracellular markers of interest.
+
+MATERIALS PROVIDED IN KIT
+================================================================================
+
+• BrdU (32.5 mM, 10 mg/mL): 5 x 1 mL sterile vials; store at ≤–70°C. Avoid multiple freeze-thaws. Vial should be opened under sterile conditions.
+• DNase I (1 mg/mL; 0.3 mg/vial): 10 x 0.3 mL vials; store at ≤–70°C. Each vial can be used to treat 10 samples. Avoid multiple freeze-thaws.
+• Anti-BrdU Antibody (clone BU20A), fluorochrome-conjugated: 1 x 100 test vial; store at 2–8°C.
+• BrdU Staining Buffer Concentrate (4X): 1 x 30 mL bottle; store at 2–8°C. This buffer contains formaldehyde. Please handle appropriately.
+• Fixation/Permeabilization Diluent: 1 x 100 mL bottle; store at 2–8°C.
+
+OTHER MATERIALS NEEDED
+================================================================================
+
+• Sterile 1X PBS
+• Flow Cytometry Staining Buffer
+• 12 x 75 mm round bottom test tubes
+• Optional:
+  - Primary antibodies (directly conjugated)
+  - Fixable Viability Dye (FVD): eFluor 450, eFluor 660, eFluor 780, eFluor 506, eFluor 520, or eFluor 455UV
+
+NOTE: The antibodies used for surface staining can be added after BrdU and Fixable Viability Dye labeling (but before fixation). Alternatively, if the antibody(s) is known to recognize a formaldehyde-fixed epitope, it can be added concurrently with the BrdU antibody.
+
+BrdU staining buffer working solution preparation
+
+Prepare fresh 1X BrdU Staining Buffer working solution by diluting BrdU Staining Buffer Concentrate (1 part) with Fixation/Permeabilization Diluent (3 parts). Mix by gentle inversion, do not vortex. You will need 1 mL of the 1X BrdU Staining Buffer working solution for each sample. Use caution and handle appropriately as the buffer contains formaldehyde.
+
+PROTOCOL
+================================================================================
+
+Step 1: In vitro labeling of 10⁵ to 10⁸ dividing cells with 10 µM BrdU for 45 minutes at 37°C.
+
+1. Under sterile conditions thaw BrdU on ice and dilute to a working concentration of 1 mM with sterile 1X PBS.
+2. Add 10 µM BrdU to each sample. (For example, add 10 µL of 1 mM BrdU directly to every milliliter of tissue culture medium.)
+3. Incubate your cells long enough to allow incorporation of BrdU. The timing will be dependent on your culture conditions (e.g., stimulants used) and the proliferation kinetics of your cells. Therefore, the incubation time will need to be determined empirically. After the incubation, harvest the cells.
+4. Wash cells by adding 2 mL of Flow Cytometry Staining Buffer (or azide-free PBS if proceeding to Step 2) and then centrifuge at 300–400 x g for 5 minutes at room temperature. Discard the supernatant.
+
+Step 2: [Optional] Stain with fixable viability dye (FVD) to label dead cells before fixation.
+
+Note: Allow the vial of Fixable Viability Dye to equilibrate to room temperature before opening. The dye must be used with azide-free PBS. For consistent staining of cells in tubes, do not stain in less than 0.5 mL. Please refer to our website Best Protocols (Viability Staining Protocol, Protocol C: Staining Dead Cells with Fixable Viability eFluor Dyes) for additional information. (Proceed to Step 3 if a FVD will not be used.)
+
+1. Wash cells one additional time with 2 mL of azide-free PBS, as described in step 4 above.
+2. Resuspend cells at 1–10 x 10⁶ cells/mL in azide-free PBS.
+3. Add 1 μL of Fixable Viability Dye per 1 mL of cells and vortex immediately.
+4. Incubate for 30 minutes at 2–8°C in the dark.
+5. Wash cells 1–2 times with Flow Cytometry Staining Buffer, as described in step 4 above.
+6. Resuspend cells at 1–10 x 10⁶ cells/mL in Flow Cytometry Staining Buffer.
+
+Step 3: [Optional] Stain cell surface antigen(s).
+
+Note: For additional information, please refer to Staining Cell Surface Antigens for Flow Cytometry. Proceed to Step 4 if cell surface antigens will not be examined or if the antibody(s) is known to recognize a formaldehyde-fixed epitope.
+
+1. Aliquot 50 μL of cell suspension to each tube or well. The cell number should be determined empirically but can range from 10⁵ to 10⁸ cells/test per tube.
+2. Add the recommended amount (refer to the Technical Data Sheet for each product) of each fluorochrome-conjugated primary antibody(s) in an appropriate volume of Flow Cytometry Staining Buffer such that the final staining volume is 100 μL. (For example, add 50 μL of an antibody mix to 50 μL of cells.) Mix gently.
+3. Incubate for at least 30 minutes at 2–8°C in the dark.
+4. Wash the cells twice with Flow Cytometry Staining Buffer, as described in step 4 under Step 1.
+
+Step 4: Fix cells and intracellularly stain with Anti-BrdU.
+
+1. Thaw DNase I solution on ice. Once thawed, prepare a working solution of DNase I by adding 300 µL of the DNase I solution to 700 µL of Flow Cytometry Staining Buffer and mix gently. Store on ice until ready for use in step 6 below.
+2. If cells were not stained in Steps 2 or 3, aliquot 100 μL of cell suspension to each tube. The cell number should be determined empirically but can range from 10⁵ to 10⁸ cells/tube.
+3. Gently resuspend the cells by pulse-vortexing once. This resuspension step is critical before the addition of the freshly prepared 1X BrdU Staining Buffer working solution.
+4. Add 1 mL of freshly prepared 1X BrdU Staining Buffer working solution and mix gently. Incubate for 15 minutes at room temperature in the dark. Incubations may go longer (up to 14 hours) but should be determined empirically for each cell type.
+5. Wash cells twice with Flow Cytometry Staining Buffer, as described in step 4 under Step 1.
+6. Add 100 µL of the DNase I working solution that was prepared in step 1 to each sample. Incubate for 1 hour at 37°C in the dark.
+7. Wash cells twice with Flow Cytometry Staining Buffer, as described in step 4 under Step 1.
+8. Add 5 µL of Anti-BrdU fluorochrome-conjugated antibody per sample. Mix and incubate for 20–30 minutes at room temperature in the dark.
+
+NOTE: Antibodies against intracellular antigens or surface antigens not stained in Step 3 may be added here. The antibodies used for surface staining at this step must recognize a fixed epitope. If an antibody only recognizes a native epitope or if this information is unknown, we recommend surface staining at Step 3.
+
+9. Wash cells twice with Flow Cytometry Staining Buffer, as described in step 4 under Step 1.
+
+Step 5: Acquire data on a flow cytometer.
+
+PRODUCT CATALOG NUMBERS
+================================================================================
+
+8811-6600-42, 8817-6600-42, 8812-6600-42, 8848-6600-42, 8846-6600-42, 00-5525-00, 00-4222-26

biomni/tool/protocols/thermofisher/BrdU and EdU Double Staining Protocol _ Thermo Fisher Scientific - US.txt

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+PROTOCOL: BrdU and EdU Double Staining Protocol
+SOURCE: Thermo Fisher Scientific - US
+URL: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/flow-cytometry-protocol/cell-proliferation/dual-pulse-labeling-of-cell-proliferation-using-edu-and-brdu-incorporation.html
+================================================================================
+
+Introduction
+The measurement of cell proliferation is fundamental to the assessment of cell health, genotoxicity, and drug efficacy. Proliferation is traditionally assessed by incubating cells with a single "pulse" of a nucleoside analog that is incorporated into DNA and detected using radioactivity, antibodies, or click chemistry. Some applications, such as drug efficacy testing, benefit from the incorporation of two different analogs at different time points (dual pulse labeling), which can further define cell cycle kinetics. With the availability of new highly specific anti-BrdU antibodies, the BrdU labeling technique can be combined with click chemistry detection for a simplified method of dual pulse labeling.
+Traditionally, the detection of cell proliferation has employed the incorporation of the thymidine analog BrdU (5-bromo-2´-deoxyuridine) during DNA synthesis, followed by detection with an anti-BrdU antibody [1–4]. This method is rapidly being replaced by the click chemistry–based Click-iT EdU assay [5,6], because unlike BrdU assays, Click-iT EdU assays are not antibody-based and therefore do not require DNA denaturation for detection of the incorporated nucleoside. Click-iT assays use a modified nucleoside, EdU (5-ethynyl-2´-deoxyuridine), that is incorporated during DNA synthesis and detected using a click reaction—a copper(I)-catalyzed reaction between an azide and an alkyne.
+
+Materials
+- Jurkat cells in complete RPMI media/10%FBS
+- BrdU (5-bromodeoxyuridin) (B23151) (10mM in DMSO)
+- EdU (5-Ethynyl-2'-deoxyuridine) (10mM in DMSO)
+- PBS pH7.2 (GIBCO(R))
+- 70% Ethanol, ice cold
+- 4M HCl
+- Phosphate/citric acid buffer, pH 7.4 (182 ml of 0.2M Na2HPO4+18ml 0.1M citric acid)
+- Antibody Diluting buffer (PBS + 0.1% Triton-X100 + 1% BSA)—prepare fresh
+- Purified anti-BrdU antibody(clone MoBU-1) (B35141) 0.1 mg/ml; use 2.5ul
+- anti-BrdU- (clone Br3) pure MD5300 lot 0903 0.4mg/ml titer use: 2.5uL
+- Goat anti-mouse IgG Alexa Fluor® 488 secondary antibody 2mg/ml, dilute 1:20 and use 2.5ul/test (0.25ug/test) (A11001)
+- Tris Buffered Saline
+- CuSo4 100mM
+- 100mM ascorbate (Freshly prepared)
+- AF647-azide 20uM DMSO A10277
+- FxCycle™ Violet 1mg/ml DI F10347 use 0.5µl/0.5ml
+
+Procedure
+1. Use Jurkat cells in complete RPMI/10%FBS media, separate into 5 flasks:
+   - Flask 1: control
+   - Flask 2: 10uM BrdU 1 hour
+   - Flask 3: 20uM EdU 1 hour
+   - Flask 4: 10µM BrdU + 20µM EdU together 1 hour
+   - Flask 5: 20µM EdU one hour then 10µM BrdU one hour
+   Incubate 37°C/5% CO2.
+
+2. After incubation, centrifuge cells, pellet and remove supernatant.
+
+3. FIX cells: Add 10ml ice cold 70% Ethanol to a 15ml tube containing the cell pellet, adding dropwise at first while vortexing, mix well.
+
+4. Store at -20°C until use, 4 hours to several months.
+
+5. Centrifuge cells, pellet & discard supernatant; vortex pellet.
+
+6. Resuspend pellet in 2.0ml 4M HCl for 20 min at room temperature.
+
+7. Add 10ml phosphate/citric acid buffer, centrifuge and decant supernatant. Filter out large clumps.
+
+8. Resuspend in PBS/0.1%Triton/1%BSA (antibody stain solution) wash & pellet.
+
+9. Repeat wash.
+
+10. Resuspend in PBS/0.1%Triton/1%BSA (antibody stain solution), adjust concentration to 1x10⁷/ml.
+
+11. Add 100µl cell suspension to designated tubes.
+
+12. Prepare click reaction and add 0.5ml to designated tubes.
+
+13. Incubate 30 minutes at room temperature protected from light.
+
+14. Wash x1 PBS/0.1%Triton/1%BSA.
+
+15. Add anti-BrdU purified antibody to designated tubes.
+
+16. Incubate for 30 minutes at room temperature protected from light.
+
+17. Wash with 3ml PBS/0.1%Triton/1%BSA (antibody stain solution) spin & pellet.
+
+18. Add Goat anti-mouse IgG Alexa Fluor 488 secondary antibody, incubate for 30 minutes at room temperature, followed by wash as in step 15.
+
+19. Resuspend pellet in 0.5ml PBS/0.1%Triton/1%BSA (antibody stain solution).
+
+20. Add 0.5µL FxCycle Violet Stain.
+
+21. Incubate for 15-30 minute at room temperature protected from light.
+
+22. Run on LSRII.
+
+23. Analyze Alexa Fluor 488 using blue laser excitation and 525/50 emission filter, Alexa Fluor 647 using red laser excitation and 660/20 emission filter, and FxCycleViolet using violet laser excitation and 450/50 emission filter.
+
+24. Gate on main population cells and collect 10,000 gated events for single color data. For triple color samples, make a singlet gate on WvA and collect 20,000 singlet events.
+
+References
+- Gratzner HG (1982) Science 218:474–475.
+- Dolbeare F, Selden JR (1994) Methods Cell Biol 41:297–316.
+- Li X, Darzynkiewicz Z (1995) Cell Prolif 28:571–579.
+- Conboy MJ, Karasov AO, Rando TA (2007) PLoS Biology 5:1120–1126.
+- Salic A, Mitchison TJ (2008) Proc Natl Acad Sci U S A 105:2415–2420.
+- Buck SB, Bradford J, Gee KR et al. (2008) Biotechniques 44:927–929.
+- Current Protocols in Cytometry Vol. 1, JP Robinson, Ed., John Wiley & Sons Inc. (2007)

biomni/tool/protocols/thermofisher/Caspase Activity Assay Protocol _ Thermo Fisher Scientific - US.txt

@@ -0,0 +1,94 @@
+PROTOCOL: Caspase Activity Assay Protocol
+SOURCE: Thermo Fisher Scientific - US
+URL: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/apoptosis-protocol.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+Caspases are a family of cysteine proteases that play essential roles in apoptosis (programmed cell death). Measuring caspase activity is a common method for detecting and quantifying apoptosis. This protocol describes methods for detecting caspase activity using fluorescent substrates in both live and fixed cells.
+
+APPLICATIONS
+================================================================================
+
+This protocol can be used for:
+• Detecting apoptosis in cell cultures
+• Quantifying caspase-3/7, caspase-8, or caspase-9 activity
+• Screening for apoptosis-inducing compounds
+• Time-course studies of apoptosis
+
+MATERIALS REQUIRED
+================================================================================
+
+• Cells in culture
+• CellEvent Caspase-3/7 Green Detection Reagent or similar caspase substrate
+• Cell culture medium
+• Phosphate-buffered saline (PBS)
+• Fluorescence microscope or microplate reader
+• Optional: NucBlue Live ReadyProbes Reagent for nuclear counterstaining
+
+PROTOCOL
+================================================================================
+
+For live cell imaging (CellEvent Caspase-3/7 Green Detection Reagent)
+
+1. Prepare cells in appropriate culture vessel (coverslips, multiwell plates, etc.).
+
+2. Treat cells with apoptosis-inducing agent or control treatment.
+
+3. Prepare CellEvent Caspase-3/7 Green Detection Reagent:
+   - Add 100 µL of PBS to the dry powder to make a 100X stock solution.
+   - Dilute 1:100 in cell culture medium to make working solution (final concentration typically 2-5 µM).
+
+4. Remove cell culture medium from cells.
+
+5. Add CellEvent Caspase-3/7 Green Detection Reagent working solution to cells.
+
+6. Incubate for 30-60 minutes at 37°C, protected from light.
+
+7. Optional: Add NucBlue Live ReadyProbes Reagent (1 drop per mL) for nuclear counterstaining.
+
+8. Wash cells once with PBS or fresh culture medium.
+
+9. Image cells immediately using a fluorescence microscope with appropriate filter set (FITC/GFP filter for green fluorescence).
+
+For microplate reader detection
+
+1. Seed cells in 96-well microplate.
+
+2. Treat cells with apoptosis-inducing agent or control.
+
+3. Prepare CellEvent Caspase-3/7 Green Detection Reagent as described above.
+
+4. Add reagent to cells and incubate as described above.
+
+5. Read fluorescence using a microplate reader:
+   - Excitation: ~502 nm
+   - Emission: ~530 nm
+
+6. Calculate caspase activity relative to controls.
+
+For flow cytometry
+
+1. Prepare cell suspension.
+
+2. Treat cells with apoptosis-inducing agent.
+
+3. Prepare CellEvent Caspase-3/7 Green Detection Reagent working solution.
+
+4. Add reagent to cells and incubate for 30-60 minutes at 37°C.
+
+5. Wash cells with PBS.
+
+6. Analyze by flow cytometry using FITC/GFP channel.
+
+PROTOCOL TIPS
+================================================================================
+
+• Include positive control (cells treated with known apoptosis inducer) and negative control (untreated cells)
+• Optimize incubation time based on your cell type and treatment
+• Protect from light during incubation to prevent photobleaching
+• For time-course studies, image at multiple time points
+• Caspase activity typically peaks 2-6 hours after apoptosis induction
+• Some caspase substrates are cell-permeant and work in live cells; others require cell fixation
+• For fixed cells, follow manufacturer's instructions for fixation and permeabilization steps

biomni/tool/protocols/thermofisher/Cell Freezing Protocols _ Thermo Fisher Scientific - US.txt

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+PROTOCOL: Cell Freezing Protocols
+SOURCE: https://www.thermofisher.com/us/en/home/references/gibco-cell-culture-basics/cell-culture-protocols/freezing-cells.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+After subculturing your adherent or suspension cells, you may need to freeze any cells not required for immediate experiments. This cell freezing protocol outlines the necessary preparations and materials to order before beginning the freezing process. It also provides step-by-step instructions that you can follow during the procedure or refer to for future use.
+
+Cryopreservation of cells, or the freezing of cells, is a technique used to store living cells and tissues at extremely low temperatures to preserve their structure over an indefinite period. By lowering the temperature to below –130°C, extracellular ice forms, decreasing kinetic and molecular activity within the cells and slowing biological aging.
+
+Why freeze your cells?
+
+• Freezing cells is crucial for maintaining an adequate cell stock
+• Cell lines are valuable resources whose replacement is both expensive and time-consuming
+• Cryopreservation of finite cell lines prevents aging and transformation
+• Freezing cells in continuous culture can prevent:
+  - genetic drift
+  - senescence
+  - microbial contamination
+  - equipment failure
+
+When a small surplus of cells becomes available from subculturing, a highly effective method is to freeze them as a seed stock, keeping them protected and not available for general laboratory use. Working stocks can be prepared and replenished from these frozen seed stocks. If the seed stocks become depleted, cryopreserved working stocks can serve as a source for preparing fresh seed stock with minimal generation increase from the initial freezing.
+
+MATERIALS REQUIRED
+================================================================================
+
+We suggest using the following materials for the cell freezing protocol:
+
+• Log-phase cultured cells
+• Complete growth medium: basal medium, serum, and supplements pre-warmed to 37°C
+• Cryoprotective agent such as DMSO (use a bottle set aside for cell culture; open only in a laminar flow hood) or a freezing medium such as Synth-a-Freeze Cryopreservation Medium or Recovery Cell Culture Freezing Medium
+• Disposable, sterile 15 mL or 50 mL conical tubes
+• Reagents and equipment to determine viable and total cell counts (e.g., Countess Automated Cell Counter, or hemocytometer, cell counter, and Trypan Blue)
+• Sterile cryogenic storage vials (i.e., cryovials)
+• Controlled rate freezing apparatus or isopropanol chamber
+• Liquid nitrogen storage container
+
+For freezing adherent cells, in addition to the above materials, you will need:
+
+• Balanced salt solution such as Gibco Dulbecco's Phosphate Buffered Saline (DPBS), containing no calcium, magnesium, or phenol red
+• Dissociation reagent such as trypsin or Gibco TrypLE Express, without phenol red
+
+Catalog numbers: J66650.AK, 339650, 5000-0020, 5100-0050, 14190250, 25200072, 12604039, 15250061, 12648010, R00550, A49862
+
+MEDIUM FOR FREEZING CELLS
+================================================================================
+
+Cryoprotective agents reduce the freezing point of the medium and slow the cooling rate, greatly reducing the risk of ice crystal formation, which can damage cells and cause cell death. Media for cryopreservation typically includes a base medium, a cryopreservative, and a protein source, but we recommend that you closely follow your cell line and product-specific instructions to determine the best cryopreservation media for your experiment. When included, the cryopreservative and protein protect the cells from the stress of the freeze-thaw process. The cells are then frozen in liquid nitrogen for preservation.
+
+For media with serum, the constituents may be:
+
+• complete medium containing 10% glycerol
+• complete medium containing 10% DMSO (dimethyl sulfoxide)
+• 50% cell-conditioned medium with 50% fresh medium with 10% glycerol or 10% DMSO
+
+Safety Note: DMSO solutions are known to facilitate the entry process of organic molecules into tissues. You should always handle reagents containing DMSO using equipment and practices appropriate for the hazards posed by such materials and dispose of the reagents in compliance with local regulations.
+
+A serum-free medium has generally low or no protein; but you can still use it as a base for a cryopreservative medium in the following formulations:
+
+• 50% cell-conditioned serum-free medium with 50% fresh serum-free medium containing 7.5% DMSO
+• fresh serum-free medium containing 7.5% DMSO and 10% cell culture–grade BSA
+
+You may also use a specially formulated complete cryopreservation medium such as:
+
+• Gibco Recovery Cell Culture Freezing is a ready-to-use complete cryopreservation medium for mammalian cell cultures with an optimized ratio of fetal bovine serum to bovine serum for improved cell viability and cell recovery after thawing.
+• Gibco Synth-a-Freeze Cryopreservation Medium is a chemically defined, protein-free, sterile cryopreservation medium with 10% DMSO that is suitable for the cryopreservation of many stem and primary cell types apart from melanocytes.
+
+PROTOCOL
+================================================================================
+
+The proper way to freeze cells and cryopreserve your cell lines for future use requires diligent attention to passaging procedures and storage requirements. Remember to always wear personal protective equipment and use proper sterile technique when freezing cells.
+
+The general freezing method is the same for adherent and suspension cells, except that adherent cells need to be removed from the culture plates before starting the freezing procedure. The following cell freezing protocol is a general procedure for cryopreserving cultured cells. Optimal freezing conditions depend on the cell line in use—for detailed protocols, always refer to the cell-specific product insert.
+
+Prior to cryopreservation, cells should be characterized and checked for contamination. Freeze cells in log phase, at a high concentration of at least 90% viability, and at as low a passage number as possible, as this will lead to the best outcomes when your stock is thawed.
+
+1. Prepare freezing medium and store at 2° to 8°C until use.
+
+2. For adherent cells, gently detach cells from the tissue culture vessel following the procedure used during subculture. Detach as gently as possible to minimize damage to the cells.
+
+3. Resuspend the detached cells in the complete growth medium.
+
+4. Find the total number of cells and percent viability using a hemocytometer or the Countess Automated Cell Counter and Trypan Blue exclusion.
+
+5. According to the desired viable cell density, calculate the required volume of freezing medium.
+
+6. Centrifuge the cell suspension at approximately 100–400 × g for 5 to 10 minutes (centrifugation speed and duration varies depending on the cell type). Using a pipette, aseptically withdraw the supernatant to the smallest volume without disturbing the cell pellet and discard.
+
+7. Resuspend the cell pellet in cold freezing medium at the recommended viable cell density for the specific cell type.
+
+8. Dispense aliquots of the cell suspension into sterile cryogenic storage vials. As you aliquot them, gently and often mix the cells to maintain a homogeneous cell suspension.
+
+9. Freeze the cells slowly by reducing the temperature at approximately 1°C per minute using a controlled rate cryo-freezer or a cryo-freezing container such as "Mr. Frosty," available from Thermo Scientific Nalgene labware (Nalgene Nunc). Alternatively, place the cryovials containing the cells in an isopropanol chamber and store them at –80°C overnight.
+
+10. Transfer frozen cells to liquid nitrogen and store them in the gas phase above the liquid nitrogen below –135°C.
+
+Safety Note: Biohazardous materials must be stored in the gas phase above the liquid nitrogen. Storing the sealed cryovials in the gas phase reduces the risk of explosion. If you are using liquid-phase storage, be aware of the explosion hazard with both glass and plastic cryovials and always wear a face shield or goggles.
+
+REFERENCES
+================================================================================
+
+Freshney, R. (1987) Culture of Animal Cells: A Manual of Basic Technique, p. 220, Alan R. Liss, Inc., New York.
+
+Mohamed, H. M., Sundar, P., Ridwan, N. A. A., Cheong, A. J., Mohamad Salleh, N. A., Sulaiman, N., Mh Busra, F., & Maarof, M. (2024). Optimisation of cryopreservation conditions, including storage duration and revival methods, for the viability of human primary cells. BMC Molecular & Cell Biology, 25(1), 1–14. https://doi.org/10.1186/s12860-024-00516-6.

biomni/tool/protocols/thermofisher/CellTrace CFSE Cell Proliferation Kit Protocol _ Thermo Fisher Scientific - US.txt

@@ -0,0 +1,94 @@
+PROTOCOL: CellTrace CFSE Cell Proliferation Kit Protocol
+SOURCE: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/protocols/celltrace-cfse-cell-proliferation-kit-protocol.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+The CellTrace CFSE Cell Proliferation Kit provides a robust method for evaluating cell proliferation through CFSE staining. CFSE, or carboxyfluorescein succinimidyl ester, is a fluorescent dye used in CFSE proliferation assays to monitor distinct generations of proliferating cells. The CFSE staining protocol involves covalently labeling live cells with this bright and stable dye, which dilutes evenly among daughter cells during cell division. Flow cytometric visualization then allows researchers to observe each generation as a separate peak on a flow cytometry histogram, offering precise insights into cell proliferation dynamics. The CellTrace CFSE protocol is essential for studies requiring accurate measurement of cell proliferation and is widely used in various research applications to evaluate cell division and growth.
+
+APPLICATIONS
+================================================================================
+
+This protocol can be used for:
+• Detecting cell proliferation using flow cytometry
+
+This protocol should not be used for:
+• Fluorescence microscopy or microplate readers
+
+MATERIALS REQUIRED
+================================================================================
+
+You will need the following for this protocol:
+
+• 20 mL of heparinized peripheral whole blood
+• CellTrace CFSE Cell Proliferation Kit
+• OpTmizer T Cell Expansion SFM
+• Penicillin-Streptomycin-Glutamine (100X)
+• PBS
+• Ficoll-Paque Plus
+• Cell counter such as the Countess 3 FL
+• Dynabeads Human T-Activator CD3/CD28
+
+PROTOCOL
+================================================================================
+
+Culture medium preparation
+
+To 1 L Gibco OpTmizer T Cell Expansion SFM, add the following:
+• 26 mL of T Cell Expansion Supplement
+• 10 mL of Penicillin-Streptomycin-Glutamine
+
+Complete medium is stable for 4 weeks when stored at 2–8°C in the dark.
+
+Mononuclear cell isolation from whole blood
+
+1. Dilute 10 mL whole blood in 10 mL PBS and mix well.
+2. Add 15 mL Invitrogen Ficoll-Paque Plus to a 50 mL centrifuge tube and gently layer 20 mL diluted whole blood on top.
+3. Centrifuge for 30 minutes at 400 x g.
+4. Carefully remove lymphocyte layer and transfer to a new tube.
+5. Resuspend cells in 25 mL DPBS buffer in a 50 mL conical tube.
+6. Centrifuge for 5 minutes at 300 x g, pour off supernatant, and resuspend in 25 mL DPBS.
+7. Repeat wash step and resuspend in 10 mL DPBS.
+8. Count cells on the Invitrogen Countess Automated Counter or by another method; adjust concentration to 10⁶ cells/mL.
+
+Cell staining
+
+1. Add 18 µL DMSO to a vial of CellTrace CFSE staining solution.
+2. Dilute this stock solution into 20 mL of PBS (warmed to 37°C) for a 5 µM staining solution.
+3. Add 10 mL of cells to a 50 mL centrifuge tube.
+4. Centrifuge cells for 5 minutes at 300 x g and carefully pour off supernatant.
+5. Resuspend cells in 10 mL of CellTrace CFSE staining solution.
+6. Incubate cells for 20 minutes in a 37°C water bath.
+7. Add 40 mL OpTmizer T Cell Expansion SFM to the cells to absorb any unbound dye.
+8. Incubate cells for 5 minutes.
+9. Centrifuge cells for 5 minutes at 300 x g and resuspend the cell pellet in pre-warmed OpTmizer T Cell Expansion SFM.
+
+Stimulation and analysis
+
+1. Distribute aliquots of stained cells into culture plates or flasks.
+2. Stimulate with 50 µL Invitrogen Dynabeads Human T-Activator CD3/CD28 per 1 mL of cells, or other stimulus.
+3. Incubate for desired length of time under growth conditions.
+4. Harvest cells and stain for other markers if appropriate.
+5. Analyze using a flow cytometer with 488 nm excitation and emission filters appropriate for fluorescein.
+
+SPECTRAL INFORMATION AND STORAGE
+================================================================================
+
+CellTrace CFSE:
+• Excitation/Emission (in nm): 492/517
+• Standard filter set: Invitrogen Alexa Fluor 488
+• Storage conditions: ≤–20°C
+
+PROTOCOL TIPS
+================================================================================
+
+• Reserve 1 mL of cells for unstained control and 1 mL of cells for a stained, but unstimulated control
+• Dynabeads stimulation typically results in T cell division every 18–20 hr
+• Analyze as many cells as possible from each sample
+• Use a viability dye and gate on live cells
+
+PRODUCT CATALOG NUMBERS
+================================================================================
+
+C34554, A1048501, 10378016, 10010023, AMQAF2000, 11161D

biomni/tool/protocols/thermofisher/CellTrace Far Red Cell Proliferation Kit Protocol _ Thermo Fisher Scientific - US.txt

@@ -0,0 +1,101 @@
+PROTOCOL: CellTrace Far Red Cell Proliferation Kit Protocol
+SOURCE: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/protocols/celltrace-far-red-cell-proliferation-kit-protocol.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+Flow cytometric visualization of cell generations
+
+The CellTrace Far Red kit is used to monitor distinct generations of proliferating cells by dye dilution. Live cells are covalently labeled with a very bright, stable dye. Every generation of cells appears as a different peak on a flow cytometry histogram.
+
+APPLICATIONS
+================================================================================
+
+This protocol can be used for:
+• Detecting cell proliferation using flow cytometry
+
+This protocol should not be used for:
+• Fluorescence microscopy or microplate readers
+
+MATERIALS REQUIRED
+================================================================================
+
+You will need the following for this protocol:
+
+• 20 mL of heparinized peripheral whole blood
+• CellTrace Far Red Cell Proliferation Kit (Cat. No. C34564)
+• OpTmizer T Cell Expansion SFM (Cat. No. A10485-01)
+• Penicillin-Streptomycin-Glutamine (100X) (Cat. No. 10378-016)
+• PBS (Cat. No. 10010-023)
+• GE Ficoll-Paque™ Plus (Fisher Cat. No. 45-001-750)
+• Cell counter such as the Countess II FL (Cat. No. AMQAF1000)
+• Dynabeads Human T-Activator CD3/CD28 (Cat. No. 11161D)
+
+PROTOCOL
+================================================================================
+
+Culture medium preparation
+
+To 1 L OpTmizer T Cell Expansion SFM, add the following:
+• 26 mL of T Cell Expansion Supplement (supplied in Cat. No. A10485-01)
+• 10 mL of Penicillin-Streptomycin-Glutamine
+
+Complete medium is stable for 4 weeks when stored at 2–8°C in the dark.
+
+Mononuclear cell isolation from whole blood
+
+1. Dilute 20 mL whole blood in 20 mL PBS and mix well.
+2. Add 15 mL Ficoll-Paque™ Plus to a 50 mL centrifuge tube and gently layer 20 mL diluted whole blood on top.
+3. Centrifuge for 30 minutes at 400 x g.
+4. Carefully remove lymphocyte layer and transfer to a new tube.
+5. Resuspend cells in 25 mL DPBS buffer in a 50 mL conical tube.
+6. Centrifuge for 5 minutes at 300 x g, pour off supernatant, and resuspend in 25 mL DPBS.
+7. Repeat wash step and resuspend in 10 mL DPBS.
+8. Count cells on the Countess Automated Counter or by another method; adjust concentration to 10⁶ cells/mL.
+
+Cell staining
+
+1. Add 20 µL DMSO to a vial of CellTrace Far Red staining solution.
+2. Dilute this stock solution into 20 mL of PBS (warmed to 37°C) for a 1 µM staining solution.
+3. Add 10 mL of cells to a 50 mL centrifuge tube.
+4. Centrifuge cells for 5 minutes at 300 x g and carefully pour off supernatant.
+5. Resuspend cells in 10 mL of CellTrace Far Red staining solution.
+6. Incubate cells for 20 minutes in a 37°C water bath.
+7. Add 40 mL OpTmizer T Cell Expansion SFM to the cells to absorb any unbound dye.
+8. Incubate cells for 5 minutes.
+9. Centrifuge cells for 5 minutes at 300 x g and resuspend the cell pellet in pre-warmed OpTmizer T Cell Expansion SFM.
+
+Stimulation and analysis
+
+1. Distribute aliquots of stained cells into culture plates or flasks.
+2. Stimulate with 50 µL Dynabeads Human T-Activator CD3/CD28 per 1 mL of cells, or other stimulus.
+3. Incubate for desired length of time under growth conditions.
+4. Harvest cells and stain for other markers if appropriate.
+5. Analyze using a flow cytometer with 633/635 nm excitation and a 660/20 bandpass emission filter.
+
+SPECTRAL INFORMATION AND STORAGE
+================================================================================
+
+CellTrace Far Red:
+• Excitation/Emission (in nm): 630/661
+• Standard filter set: Alexa Fluor 647
+• Storage conditions: ≤–20°C
+
+PROTOCOL TIPS
+================================================================================
+
+• Reserve 1 mL of cells for unstained control and 1 mL of cells for a stained, but unstimulated control
+• Dynabeads stimulation typically results in T cell division every 18–20 hr
+• Analyze as many cells as possible from each sample
+• Use a viability dye and gate on live cells
+
+EXAMPLE APPLICATION
+================================================================================
+
+Human T lymphocytes stained with CellTrace Far Red Cell Proliferation reagent and stimulated in culture for 5 days. The discrete peaks represent successive generations of live cells. The unstimulated parent generation is indicated in purple. Analysis was completed using an Attune Acoustic Focusing Cytometer with 638 nm excitation and a 660/20 nm bandpass emission filter.
+
+PRODUCT CATALOG NUMBER
+================================================================================
+
+C34564

biomni/tool/protocols/thermofisher/CellTrace Violet Cell Proliferation Kit Protocol _ Thermo Fisher Scientific - US.txt

@@ -0,0 +1,94 @@
+PROTOCOL: CellTrace Violet Cell Proliferation Kit Protocol
+SOURCE: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/protocols/celltrace-violet-cell-proliferation-kit-protocol.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+The CellTrace Violet (CTV) Cell Proliferation Kit is a powerful tool used for monitoring cell proliferation through dye dilution. In this CellTrace Violet protocol, live cells are covalently labeled with a very bright and stable dye which is retained within the cells and evenly distributed to daughter cells during cell division allowing researchers to track distinct generations of proliferating cells. This process, known as CTV staining, results in each cell generation appearing as a different peak on a flow cytometry histogram, facilitating precise evaluation of cell proliferation. These methods provide valuable insights into cellular behaviors and responses, making the CellTrace Violet kit a key tool for flow cytometric visualization of cell generations.
+
+APPLICATIONS
+================================================================================
+
+This protocol can be used for:
+• Detecting cell proliferation using flow cytometry
+
+This protocol should not be used for:
+• Fluorescence microscopy or microplate readers
+
+MATERIALS REQUIRED
+================================================================================
+
+You will need the following for this protocol:
+
+• 10 mL of heparinized peripheral whole blood
+• CellTrace Violet Cell Proliferation Kit
+• OpTmizer T Cell Expansion SFM
+• Penicillin-Streptomycin-Glutamine (100X)
+• PBS
+• Ficoll-Paque Plus
+• Cell counter such as the Countess 3 FL
+• Dynabeads Human T-Activator CD3/CD28
+
+PROTOCOL
+================================================================================
+
+Culture medium preparation
+
+To 1 L OpTmizer T Cell Expansion SFM, add the following:
+• 26 mL of T Cell Expansion Supplement
+• 10 mL of Penicillin-Streptomycin-Glutamine
+
+Complete medium is stable for 4 weeks when stored at 2–8°C in the dark.
+
+Mononuclear cell isolation from whole blood
+
+1. Dilute 10 mL whole blood in 10 mL PBS and mix well.
+2. Add 15 mL Ficoll-Paque Plus to a 50 mL centrifuge tube and gently layer 20 mL diluted whole blood on top.
+3. Centrifuge for 30 minutes at 400 x g.
+4. Carefully remove lymphocyte layer and transfer to a new tube.
+5. Resuspend cells in 25 mL DPBS buffer in a 50 mL conical tube.
+6. Centrifuge for 5 minutes at 300 x g, pour off supernatant, and resuspend in 25 mL DPBS.
+7. Repeat wash step and resuspend in 10 mL DPBS.
+8. Count cells on the Countess Automated Counter or by another method; adjust concentration to 10⁶ cells/mL.
+
+Cell staining
+
+1. Add 20 µL DMSO to a vial of CellTrace Violet staining solution.
+2. Dilute this stock solution into 20 mL of PBS (warmed to 37°C) for a 5 µM staining solution.
+3. Add 10 mL of cells to a 50 mL centrifuge tube.
+4. Centrifuge cells for 5 minutes at 300 x g and carefully pour off supernatant.
+5. Resuspend cells in 10 mL of CellTrace Violet staining solution.
+6. Incubate cells for 20 minutes in a 37°C water bath.
+7. Add 40 mL OpTmizer T Cell Expansion SFM to the cells to absorb any unbound dye.
+8. Incubate cells for 5 minutes.
+9. Centrifuge cells for 5 minutes at 300 x g and resuspend the cell pellet in pre-warmed OpTmizer T Cell Expansion SFM.
+
+Stimulation and analysis
+
+1. Distribute aliquots of stained cells into culture plates or flasks.
+2. Stimulate with 50 µL Dynabeads Human T-Activator CD3/CD28 per 1 mL of cells, or other stimulus.
+3. Incubate for desired length of time under growth conditions.
+4. Harvest cells and stain for other markers if appropriate.
+5. Analyze using a flow cytometer with 405 nm excitation and a 450/40 bandpass emission filter.
+
+SPECTRAL INFORMATION AND STORAGE
+================================================================================
+
+CellTrace Violet:
+• Excitation/Emission (in nm): 405/450
+• Standard filter set: Pacific Blue
+• Storage conditions: ≤–20°C
+
+PROTOCOL TIPS
+================================================================================
+
+• Reserve 1 mL of cells for unstained control and 1 mL of cells for a stained, but unstimulated control
+• Dynabeads stimulation typically results in T-cell division every 18–20 hr
+• Analyze as many cells as possible from each sample
+• Use a viability dye and gate on live cells
+
+PRODUCT CATALOG NUMBERS
+================================================================================
+
+C34557, A1048501, 10378016, 10010023, AMQAF2000, 11161D

biomni/tool/protocols/thermofisher/Click-iT EdU Protocol for Flow Cytometry _ Thermo Fisher Scientific - US.txt

@@ -0,0 +1,146 @@
+PROTOCOL: Click-iT EdU Protocol for Flow Cytometry
+SOURCE: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/protocols/click-it-edu-flow-cytometry-protocol.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+The Click-iT EdU Flow Cytometry Assay Kits provide a fast and simple method for detecting and quantifying cell proliferation using flow cytometry. EdU (5-ethynyl-2'-deoxyuridine) is a thymidine analog that is incorporated into DNA during active DNA synthesis. The Click-iT reaction uses copper-catalyzed azide-alkyne cycloaddition (CuAAC) chemistry to attach a fluorescent azide to the incorporated EdU, allowing for detection and quantification of proliferating cells.
+
+MATERIALS PROVIDED IN KIT
+================================================================================
+
+• Component A: EdU (5-ethynyl-2'-deoxyuridine), 40 mg
+• Component B: Fluorescent azide (Pacific Blue azide, Alexa Fluor 647 azide, or Alexa Fluor 488 azide)
+• Component C: DMSO, 4 mL
+• Component D: Click-iT fixative (contains paraformaldehyde)
+• Component E: Click-iT saponin-based permeabilization and wash reagent (10X), 50 mL
+• Component F: CuSO₄ solution, 1 mL
+• Component G: Click-iT EdU buffer additive
+
+Number of assays: Sufficient material is supplied for 50 reactions, based on the protocol below.
+
+Approximate fluorescence excitation/emission maxima:
+• Pacific Blue azide: 410/455 nm
+• Alexa Fluor 647 azide: 650/670 nm
+• Alexa Fluor 488 azide: 495/519 nm
+
+MATERIALS REQUIRED BUT NOT PROVIDED
+================================================================================
+
+• 1% bovine serum albumin (BSA) in phosphate buffered saline (PBS), pH 7.1–pH 7.4
+• Buffered saline solution, such as PBS, D-PBS, or TBS
+• Deionized water or 18 MΩ purified water
+• 12 × 75-mm tubes, or other flow cytometry tubes
+
+CAUTIONS
+================================================================================
+
+• DMSO (Component C), provided as a solvent in this kit, is known to facilitate the entry of organic molecules into tissues. Handle reagents containing DMSO using equipment and practices appropriate for the hazards posed by such materials. Dispose of the reagents in compliance with all pertaining local regulations.
+• Click-iT fixative (Component D) contains paraformaldehyde, which is harmful. Use with appropriate precautions.
+• Click-iT saponin-based permeabilization and wash reagent (Component E) contains sodium azide, which yields highly toxic hydrazoic acid under acidic conditions. Dilute azide compounds in running water before discarding to avoid accumulation of potentially explosive deposits in plumbing.
+
+PROTOCOL
+================================================================================
+
+Preparing reagents
+
+1. Allow vials to warm to room temperature before opening.
+2. To prepare a 10 mM solution of EdU, add 4 mL of DMSO (Component C) or aqueous solution (PBS) to Component A and mix well. After use, store any remaining stock solution at ≤–20°C. When stored as directed, the stock solution is stable for up to 1 year.
+3. To prepare a working solution of Pacific Blue azide, Alexa Fluor 647 azide, or Alexa Fluor 488 azide, add 130 μL of DMSO to Component B and mix well. After use, store any remaining working solution at ≤–20°C. When stored as directed, this working solution is stable for up to 1 year.
+4. To prepare 500 mL of 1X Click-iT saponin-based permeabilization and wash reagent, add 50 mL of Component E to 450 mL of 1% BSA in PBS. Smaller amounts can be prepared by diluting a volume of Component E 1:10 with 1% BSA in PBS. After use, store any remaining solutions at 2–6˚C. When stored as directed, the 1X solution is stable for 6 months and the 10X solution is stable for 12 months after receipt.
+
+Note: Component E contains sodium azide (see Cautions).
+
+5. To make a 10X stock solution of the Click-iT EdU buffer additive (Component G), add 2 mL of deionized water to the vial and mix until the Click-iT EdU buffer additive is fully dissolved. After use, store any remaining stock solution at ≤–20˚C. When stored as directed, the stock solution is stable for up to 1 year.
+
+Labeling cells with EdU
+
+The following protocol was developed with Jurkat cells, a human T cell line, and using an EdU concentration of 10 μM, and can be adapted for any cell type. Growth medium, cell density, cell type variations, and other factors may influence labeling. In initial experiments, we recommend testing a range of EdU concentrations to determine the optimal concentration for your cell type and experimental conditions. If currently using a BrdU based assay for cell proliferation, a similar concentration to BrdU is a good starting concentration for EdU. If using whole blood as the sample, we recommend heparin as the anticoagulant for collection.
+
+1. Suspend the cells in an appropriate tissue culture medium to obtain optimal conditions for cell growth. Disturbing the cells by temperature changes or washing prior to incubation with EdU slows the growth of the cells during incorporation.
+2. Add EdU to the culture medium at the desired final concentration and mix well. We recommend a starting concentration of 10 μM for 1–2 hours. For longer incubations, use lower concentrations. For shorter incubations, higher concentrations may be required. For a negative staining control, include cells from the same population that have not been treated with EdU.
+3. Incubate under conditions optimal for cell type for the desired length of time. Altering the amount of time the cells are exposed to EdU or subjecting the cells to pulse labeling with EdU allows the evaluation of various DNA synthesis and proliferation parameters. Effective time intervals for pulse labeling and the length of each pulse depend on the cell growth rate.
+4. Harvest cells and proceed immediately to "Staining cell-surface antigens with antibodies" if performing antibody surface labeling; otherwise continue to "Fixation and permeabilization".
+
+Staining cell-surface antigens with antibodies (optional)
+
+1. Wash cells once with 3 mL of 1% BSA in PBS, pellet cells by centrifugation, and remove supernatant.
+2. Dislodge the pellet and resuspend cells at 1 × 10⁷ cells/mL in 1% BSA in PBS.
+3. Add 100 μL of cell suspension or whole blood sample to flow tubes.
+4. Add surface antibodies and mix well.
+
+Note: Do not use PE, PE-tandem, or Qdot antibody conjugates before performing the click reaction; wait until step 6.1 for labeling with these fluorophores.
+
+5. Incubate for the recommended time and temperature, protected from light.
+6. Proceed to step 4.1 for cell fixation.
+
+Fixation and permeabilization
+
+The Click-iT saponin-based permeabilization and wash reagent can be used with whole blood or cell suspensions containing red blood cells, as well as with cell suspensions containing more than one cell type. This permeabilization and wash reagent maintains the morphological light scatter characteristics of leukocytes while lysing red blood cells.
+
+1. Wash the cells once with 3 mL of 1% BSA in PBS, pellet the cells, and remove the supernatant.
+2. Dislodge the pellet, add 100 μL of Click-iT fixative (Component D), and mix well.
+3. Incubate the cells for 15 minutes at room temperature, protected from light.
+4. Wash the cells with 3 mL of 1% BSA in PBS, pellet the cells, and remove the supernatant. Repeat the wash step if red blood cells or hemoglobin are present in the sample. Remove all residual red blood cell debris and hemoglobin before proceeding.
+5. Dislodge the cell pellet and resuspend the cells in 100 μL of 1X Click-iT saponin-based permeabilization and wash reagent (prepared in step 4 under "Preparing reagents"), and mix well. Incubate the cells for 15 minutes or proceed directly to the next protocol for click labeling.
+
+Click-iT reaction
+
+1. Prepare 1X Click-iT EdU buffer additive by diluting the 10X stock solution (prepared in step 5 under "Preparing reagents") 1:10 in deionized water.
+2. Prepare the Click-iT reaction cocktail according to Table 1.
+
+NOTE: Use the Click-iT reaction cocktail within 15 minutes of preparation.
+
+Table 1. Click-iT EdU reaction cocktails.
+
+Reaction components | 1 reaction | 2 reactions | 5 reactions | 10 reactions | 15 reactions | 30 reactions | 50 reactions
+---------------------|-----------|-------------|-------------|--------------|--------------|--------------|-------------
+PBS, D-PBS, or TBS  | 438 µL    | 875 µL      | 2.19 mL     | 4.38 mL      | 6.57 mL      | 13.2 mL      | 21.9 mL
+CuSO₄ (Component F)  | 10 µL     | 20 µL       | 50 µL       | 100 µL       | 150 µL       | 300 µL       | 500 µL
+Fluorescent dye azide | 2.5 µL   | 5 µL        | 12.5 µL     | 25 µL        | 37.5 µL      | 75 µL        | 125 µL
+Reaction Buffer Additive | 50 µL | 100 µL      | 250 µL      | 500 µL       | 750 µL       | 1.5 mL        | 2.5 mL
+Total reaction volume | 500 µL   | 1 mL        | 2.5 mL      | 5 mL         | 7.5 mL       | 15 mL         | 25 mL
+
+3. Add 0.5 mL of Click-iT reaction cocktail to each tube and mix well.
+4. Incubate the reaction mixture for 30 minutes at room temperature, protected from light.
+5. Wash the cells once with 3 mL of 1X Click-iT saponin-based permeabilization and wash reagent (prepared in step 4 under "Preparing reagents"), pellet the cells, and remove the supernatant. Dislodge the cell pellet and resuspend the cells in 100 μL of 1X Click-iT saponin-based permeabilization and wash reagent, if proceeding with intracellular antibody labeling in "Staining cells for DNA content". Otherwise, add 500 μL of 1X Click-iT saponin-based permeabilization and wash reagent and proceed to "Analysis by flow cytometry".
+
+Staining cells for DNA content (optional)
+
+1. If necessary, add Ribonuclease A to each tube and mix (see Table 2).
+2. Add the appropriate DNA stain to each tube, mix well, and incubate as recommended for each DNA stain.
+
+Table 2. Click-iT EdU compatibility with DNA content stains.
+
+DNA content stain | Click-iT EdU stain compatibility | RNase required?
+-----------------|----------------------------------|----------------
+                 | Pacific Blue | Alexa Fluor 647 | Alexa Fluor 488 |
+FxCycle Violet   | No           | Yes             | Yes             | No
+Propidium iodide (PI) | Yes      | Yes             | No*             | Yes
+SYTOX AADvanced  | Yes          | Yes             | Yes             | Yes
+FxCycle Far Red  | Yes          | No              | Yes             | Yes
+
+Analysis by flow cytometry
+
+If measuring total DNA content on a traditional flow cytometer using hydrodynamic focusing, use a low flow rate during acquisition. If using the Attune Acoustic Focusing Cytometer, all collection rates may be used without loss of signal integrity if the event rate is kept below 10,000 events per second. However, for each sample within an experiment, the same collection rate and cell concentration should be used. The fluorescent signal generated by DNA content stains is best detected with linear amplification. The fluorescent signal generated by Click-iT EdU labeling is best detected with logarithmic amplification.
+
+1. Analyze the cells using a flow cytometer.
+2. For the detection of EdU with Pacific Blue azide, use 405 nm excitation with a violet emission filter (450/50 nm or similar).
+3. For the detection of EdU with Alexa Fluor 647 azide use 633/635 nm excitation with a red emission filter (660/20 nm or similar).
+4. For the detection of EdU with Alexa Fluor 488 azide, use 488 nm excitation with a green emission filter (530/30 nm or similar).
+
+PRODUCT CATALOG NUMBERS
+================================================================================
+
+C10418, C10424, C10425, C10419, C10420, A10044, F10347, F10348, H3570, P3566, S10349, V35003, 14190144, 14190250
+
+REFERENCES
+================================================================================
+
+Chembiochem 4, 1147 (2003)
+J Am Chem Soc 125, 3192 (2003)
+Angew Chem Int Ed Engl 41, 2596 (2002)
+Angew Chem Int Ed Engl 40, 2004 (2001)
+BioTechniques 44, 927 (2008)
+Curr Protoc Cytom 55, 7.38.1 (2011)

biomni/tool/protocols/thermofisher/DAPI Protocol for Fluorescence Imaging _ Thermo Fisher Scientific - US.txt

@@ -0,0 +1,76 @@
+PROTOCOL: DAPI Protocol for Fluorescence Imaging
+SOURCE: Thermo Fisher Scientific - US
+URL: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/protocols/dapi-protocol-fluorescence-imaging.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+DAPI (4',6-diamidino-2-phenylindole) is a blue-fluorescent DNA stain that binds strongly to A-T rich regions in DNA. It is widely used for nuclear staining in fluorescence microscopy applications. DAPI is cell-impermeant and is typically used to stain fixed cells, though it can also be used to stain live cells at higher concentrations.
+
+APPLICATIONS
+================================================================================
+
+This protocol can be used for:
+• Nuclear staining in fixed cells for fluorescence microscopy
+• Cell counting and nuclear morphology analysis
+• Counterstaining in multicolor immunofluorescence experiments
+
+This protocol should not be used for:
+• Flow cytometry (use DAPI Protocol for HCA Instruments instead)
+• Live cell imaging (unless using higher concentrations and short exposure times)
+
+MATERIALS REQUIRED
+================================================================================
+
+• Fixed cells (on coverslips, slides, or in multiwell plates)
+• DAPI stain (e.g., DAPI, dihydrochloride, Cat. No. D1306, or DAPI solution)
+• Phosphate-buffered saline (PBS)
+• Mounting medium (e.g., ProLong Gold Antifade Mountant with DAPI, or add DAPI to mounting medium)
+• Fluorescence microscope with DAPI/UV filter set (excitation ~358 nm, emission ~461 nm)
+
+PROTOCOL
+================================================================================
+
+For fixed cells
+
+1. Fix and permeabilize cells according to your standard protocol.
+
+2. Wash cells 2-3 times with PBS.
+
+3. Prepare DAPI staining solution:
+   - For DAPI powder: Dissolve in water or PBS to make a 1-5 mg/mL stock solution. Store at 2-8°C, protected from light.
+   - Working solution: Dilute stock to 1-5 µg/mL in PBS or mounting medium.
+
+4. Option A: Staining before mounting
+   - Remove PBS from cells.
+   - Add DAPI working solution (1-5 µg/mL in PBS) to cover cells.
+   - Incubate for 5-10 minutes at room temperature, protected from light.
+   - Wash 2-3 times with PBS.
+   - Proceed to mounting.
+
+5. Option B: Staining in mounting medium (recommended)
+   - Remove PBS from cells.
+   - Add mounting medium containing DAPI (e.g., ProLong Gold Antifade Mountant with DAPI, or add DAPI to mounting medium at 1-5 µg/mL final concentration).
+   - Cover with coverslip and proceed to imaging.
+
+6. Image cells using a fluorescence microscope with DAPI/UV filter set.
+
+SPECTRAL INFORMATION
+================================================================================
+
+DAPI:
+• Excitation maximum: ~358 nm (UV)
+• Emission maximum: ~461 nm (blue)
+• Standard filter set: DAPI/UV
+• Compatible with: FITC, TRITC, Texas Red, Alexa Fluor dyes
+
+PROTOCOL TIPS
+================================================================================
+
+• DAPI is light-sensitive; protect from light during staining and storage
+• Optimal concentration may vary; start with 1 µg/mL and adjust as needed
+• Higher concentrations (5-10 µg/mL) may be needed for some tissue sections
+• DAPI can be used in combination with other fluorescent stains (use appropriate filter sets)
+• For live cells, use higher concentrations (5-10 µg/mL) and minimize exposure time
+• DAPI-stained samples can be stored at 2-8°C, protected from light, for several weeks

biomni/tool/protocols/thermofisher/ELISA Sample Preparation - Adherent and Suspension Cells _ Thermo Fisher Scientific - US.txt

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+PROTOCOL: ELISA Sample Preparation - Adherent and Suspension Cells
+SOURCE: Thermo Fisher Scientific - US
+URL: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/elisa-protocol/elisa-sample-preparation-protocols.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+Proper sample preparation is critical for successful ELISA experiments. This protocol describes methods for preparing samples from adherent and suspension cell cultures in a 96-well format. The prepared samples can be used directly in ELISA assays or stored for later analysis.
+
+MATERIALS REQUIRED
+================================================================================
+
+• Cells in culture (adherent or suspension)
+• Cell culture medium
+• Phosphate-buffered saline (PBS), ice-cold
+• Cell lysis buffer (e.g., RIPA buffer, NP-40 lysis buffer, or commercial cell lysis buffer)
+• Protease inhibitors (optional but recommended)
+• Phosphatase inhibitors (if analyzing phosphoproteins)
+• 96-well cell culture plate
+• Cell scraper (for adherent cells)
+• Centrifuge and microcentrifuge tubes
+• BCA or other protein quantification assay kit
+
+PROTOCOL
+================================================================================
+
+For adherent cells
+
+1. Seed cells in a 96-well plate and culture until they reach the desired confluency or treatment endpoint.
+
+2. Remove cell culture medium and wash cells once with ice-cold PBS.
+
+3. Remove PBS completely.
+
+4. Add 50-100 µL of ice-cold cell lysis buffer containing protease inhibitors (and phosphatase inhibitors if needed) to each well.
+
+5. Incubate on ice for 10-30 minutes with gentle shaking or rocking.
+
+6. Scrape cells from the bottom of wells using a cell scraper or pipette tip.
+
+7. Transfer lysate to microcentrifuge tubes.
+
+8. Centrifuge at 10,000-14,000 x g for 10-15 minutes at 4°C to pellet cell debris.
+
+9. Transfer supernatant (containing soluble proteins) to new tubes.
+
+10. Quantify protein concentration using BCA assay or similar method.
+
+11. Dilute samples to appropriate concentration in sample diluent (typically PBS or assay buffer) for ELISA.
+
+12. Store samples at -80°C if not using immediately.
+
+For suspension cells
+
+1. Culture cells in appropriate vessel until they reach desired density or treatment endpoint.
+
+2. Transfer cell suspension to centrifuge tubes.
+
+3. Centrifuge at 300-500 x g for 5 minutes at 4°C to pellet cells.
+
+4. Remove supernatant and wash cell pellet once with ice-cold PBS.
+
+5. Centrifuge again and remove PBS completely.
+
+6. Resuspend cell pellet in ice-cold cell lysis buffer containing protease inhibitors (and phosphatase inhibitors if needed).
+
+7. Use 50-100 µL lysis buffer per 10⁶ cells, or adjust volume based on cell density.
+
+8. Incubate on ice for 10-30 minutes with occasional vortexing or pipetting.
+
+9. Centrifuge at 10,000-14,000 x g for 10-15 minutes at 4°C to pellet cell debris.
+
+10. Transfer supernatant (containing soluble proteins) to new tubes.
+
+11. Quantify protein concentration using BCA assay or similar method.
+
+12. Dilute samples to appropriate concentration in sample diluent for ELISA.
+
+13. Store samples at -80°C if not using immediately.
+
+For cell culture supernatant
+
+1. Culture cells in 96-well plate until treatment endpoint.
+
+2. Collect cell culture supernatant without disturbing cells.
+
+3. Centrifuge supernatant at 300-500 x g for 5 minutes at 4°C to remove any cells or debris.
+
+4. Transfer clarified supernatant to new tubes.
+
+5. Use supernatant directly in ELISA or store at -80°C.
+
+6. Note: Supernatant samples typically do not require lysis buffer, as secreted proteins are already in solution.
+
+PROTOCOL TIPS
+================================================================================
+
+• Always work on ice to minimize protein degradation
+• Include protease inhibitors in lysis buffer to prevent protein degradation
+• For phosphoprotein analysis, include phosphatase inhibitors
+• Optimize lysis buffer volume based on cell number and protein concentration
+• Avoid excessive pipetting or vortexing, which can cause protein denaturation
+• Keep samples cold throughout the preparation process
+• Aliquot samples to avoid repeated freeze-thaw cycles
+• Include appropriate controls (untreated cells, positive controls if available)

biomni/tool/protocols/thermofisher/Enzymatic and Non-Enzymatic Cell Dissociation Protocols _ Thermo Fisher Scientific - US.txt

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+PROTOCOL: Enzymatic and Non-Enzymatic Cell Dissociation Protocols
+SOURCE: https://www.thermofisher.com/us/en/home/references/gibco-cell-culture-basics/cell-culture-protocols/cell-dissociation-protocol.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+Detachment, dissociation, and disaggregation are processes that require cellular material to be broken down from adherent conditions for study—without compromising the health of the cells or cultures involved. To do this, there are a wide range of dissociation methods and reagents available. These dissociation reagents are specially formulated with detachment agents tailored to specific cell types. Below you will find methods, protocols, and materials of the most common dissociation reagents used for gently detaching cells from a substrate or breaking down primary tissues.
+
+DETACHMENT METHODS
+================================================================================
+
+There are several detachment methods used to remove cells from substrates while maintaining the health of the cells to be harvested. Some cell lines require more gentle or assertive detachment methods, so it's imperative to understand your cell line's behavior and reaction to physical, enzymatic, or non-enzymatic dissociation methods.
+
+Table 1. Detachment methods
+
+Procedure              | Dissociation agent                                    | Applications
+-----------------------|------------------------------------------------------|--------------------------------------------------------------------------------------------------
+Shake-off              | Gentle shaking or rocking of culture vessel, or vigorous pipetting | Loosely adherent cells, mitotic cells
+Scraping               | Cell scraper                                         | Cell lines sensitive to proteases; may damage some cells
+Enzymatic dissociation | Trypsin                                              | Strongly adherent cells
+Enzymatic dissociation | Trypsin + collagenase                                | High density cultures, cultures that have formed multiple layers, especially fibroblasts
+Enzymatic dissociation | Dispase                                              | Detaching epidermal cells as confluent, intact sheets from the surface of culture dishes without dissociating the cells
+Enzymatic dissociation | Gibco TrypLE dissociation enzyme                     | Strongly adherent cells; direct substitute for trypsin; applications that require animal origin-free reagents
+Non-enzymatic dissociation | Cell dissociation buffer with salt solutions      | Lightly adherent cells, gentle, applications requiring intact cell surface proteins, not recommended for strongly adherent cells
+
+ENZYMATIC CELL DISSOCIATION
+================================================================================
+
+The following is a general procedure to rapidly remove various cell lines from the substratum while maintaining cellular integrity. This procedure is not meant to be universally applicable for all cell lines. The optimal conditions and concentrations employed for individual systems should be determined empirically. Cell viability should be routinely monitored at the time of subculturing. Cell viability should be greater than 90%.
+
+1. Remove and discard spent cell culture media.
+
+2. Wash cells using a balanced salt solution without calcium and magnesium or wash with EDTA. Add wash solution to the side of the flask opposite the cells. Rinse the cell sheet by rocking the flask for 1 to 2 min and discard wash solution.
+
+3. Add the dissociation solution of choice at 2 to 3 mL/25 cm² to the side of the flask opposite the cells. Be certain that the dissociation solution covers the cell sheet. Incubate the flasks at 37°C. Rock the flasks gently. Generally, cells are dissociated in 5 to 15 min. The actual time needed to dissociate cells will vary according to cell line. Monitor the process carefully to avoid cell damage. In addition to rocking gently, flasks of cell lines that are characteristically difficult to remove from the substratum may be tapped to expedite removal.
+
+4. When the cells are completely detached, stand the flask in the upright position to allow the cells to drain to the bottom of the flask. Add complete media to the flask. Disperse the cells by pipetting the suspension repeatedly.
+
+5. Transfer cell suspension to a 15 mL conical tube and centrifuge at 100 × g for 5–10 minutes.
+
+6. Discard supernatant and resuspend cell pellet with 2–5 mL of pre-warmed complete medium.
+
+7. Determine viable cell density and percent viability using a Countess Automated Cell Counter (similar automated or manual methods may be used).
+
+8. Seed, incubate, and subculture according to normal protocols for your cell type.
+
+TRYPLE EXPRESS ENZYMES PROTOCOL
+================================================================================
+
+TrypLE Express Enzymes are formulated to allow direct substitution into your existing protocols. The following general procedure can be used to remove various cell lines from cultureware while maintaining cellular integrity. Optimal conditions and concentrations employed for individual systems should be determined empirically.
+
+Note: TrypLE Express Enzymes may be used at ambient room temperature for many types of cells.
+
+1. Pre-warm TrypLE Express Enzymes and complete growth medium to 37°C before use. Minimize dwell time.
+
+2. Aspirate spent medium and discard.
+
+3. Wash cell monolayer with 5 mL of Dulbecco's Phosphate Buffered Saline (DPBS) without calcium and magnesium. Aspirate and discard.
+
+4. Add an appropriate volume (e.g., 5 mL in a 75 cm² flask) of TrypLE Express Enzymes to flask. Ensure complete coverage of cell monolayer with TrypLE Express Enzymes.
+
+5. Incubate at 37°C until cells have detached. Observe cell monolayer using an inverted microscope to ensure complete cell detachment from the surface of the flask. Gently tap flask to dislodge cells if necessary.
+
+6. Add 5–10 mL of pre-warmed complete medium to flask. Tilt flask in all directions to thoroughly rinse flask and transfer cell suspension to a 15 mL conical tube.
+
+7. Centrifuge at 100 × g for 5–10 minutes.
+
+8. Discard supernatant and resuspend cell pellet with 2–5 mL of pre-warmed complete medium.
+
+9. Determine viable cell density and percent viability using a Countess Automated Cell Counter (similar automated or manual methods may be used).
+
+10. Seed, incubate and subculture according to normal protocols depending on your cell type.
+
+Note: Use of soybean trypsin inhibitors are not recommended post-dissociation.
+
+ENZYMATIC DISSOCIATION REAGENTS
+================================================================================
+
+Catalog numbers: 15090046, 12604013, 12605010, 17018029, 17101015, 17104019, 17105041
+
+DISAGGREGATION AND TISSUE DISSOCIATION
+================================================================================
+
+A common method to obtain single cell suspensions from primary tissue is enzymatic disaggregation. Expose the cells to enzymes for a minimal amount of time to preserve maximum viability. The following procedures disaggregate whole tissue to obtain a high yield of viable cells.
+
+TRYPSIN PRIMARY TISSUE PROTOCOL
+================================================================================
+
+1. After dissecting off unusable tissue, mince the remaining tissue into 3 to 4 mm pieces with a sterile scalpel or scissors. Wash the tissue pieces by resuspending in a balanced salt solution without calcium and magnesium. Allow the tissue pieces to settle and remove the supernatant. Repeat the wash 2 or 3 times.
+
+2. Place the container with the tissue pieces on ice and remove any remaining supernatant. Add 0.25% trypsin in a balanced salt solution without calcium or magnesium (1 mL of trypsin for every 100 mg of tissue).
+
+3. Incubate at 4°C for 6 to 18 h to maximize penetration of the enzyme with little trypsin activity.
+
+4. Decant and discard the trypsin from the tissue pieces. Incubate the tissue pieces with residual trypsin at 37°C for 20 to 30 min.
+
+5. Add warm, complete media to the tissue pieces and gently disperse the tissue by pipetting. If using a serum-free medium, add soybean trypsin inhibitor.
+
+6. Filter the cell suspension through sterile, stainless-steel mesh (100 to 200 µM) to completely disperse any remaining tissue. Count and seed the cells for culture.
+
+COLLAGENASE PRIMARY TISSUE PROTOCOL
+================================================================================
+
+1. Mince tissue into 3–4 mm pieces with a sterile scalpel or scissors.
+
+2. Wash the tissue pieces several times with HBSS containing calcium and magnesium.
+
+3. Add sufficient HBSS with calcium and magnesium to submerge tissue. Add collagenase to 50–200 U/mL.
+
+4. Incubate at 37°C for 4–18 hours. Increased efficiency is obtained using a rocker platform and supplementing the digest with 3 mM CaCl₂.
+
+5. Disperse cells by passing through a sterile stainless steel or nylon mesh. Remaining tissue fragments may be disaggregated by addition to fresh collagenase solution and further incubation at 37°C.
+
+6. Wash dispersed cells several times by centrifugation in HBSS w/o collagenase.
+
+7. Resuspend cell pellet, after the final wash step, in culture medium. Determine viable cell density using a Countess Automated Cell Counter (alternate automated or manual methods may be used).
+
+8. Seed cells into culture vessels containing appropriate media.
+
+DISPASE PRIMARY TISSUE PROTOCOL
+================================================================================
+
+1. Mince tissue into 3–4 mm pieces with a sterile scalpel or scissors.
+
+2. Wash the tissue pieces several times in sterile DPBS without calcium and magnesium.
+
+3. Submerge tissue fragments in Dispase solution (0.6–2.4 U/mL) and incubate at 37°C. More efficient dissociation of tissue can be obtained by mixing the Dispase solution at 0.3–0.6 U/mL with collagenase (60–100 U/mL).
+
+4. Stir slowly at 37°C until the tissue is sufficiently dissolved. For compact tissues, we recommend incubating for 1 hour. Cells will not be adversely affected even after several hours in Dispase solution.
+
+5. If necessary, separate the dispersed cells from residual tissue by passing the mixture through a sterile stainless steel or nylon mesh or simply decant the cells after larger fragments have settled. Fresh Dispase solution may be added to the remaining tissue fragments if further disaggregation is required.
+
+6. Pellet cells by centrifugation and decant the enzyme solution.
+
+7. Resuspend the cell pellet in appropriate culture medium. Determine viable cell density using a Countess Automated Cell Counter (alternate automated or manual methods may be used).
+
+8. Seed cells into culture vessels containing appropriate culture medium and incubate under predetermined conditions.
+
+ENZYMATIC TISSUE DISSOCIATION REAGENTS
+================================================================================
+
+Catalog numbers: 15400054, 15050065, 15050057, 12604013, 12605028, 12605010, 17100017, 17018029, 17101015, 17104019, 17105041, 14190144
+
+NON-ENZYMATIC CELL DISSOCIATION
+================================================================================
+
+The following is a general procedure to rapidly remove various cell lines from the substratum while maintaining cellular integrity. This procedure is not meant to be universally applicable for all cell lines. The optimal conditions and concentrations employed for individual systems should be determined empirically. Cell viability should be routinely monitored at the time of subculturing. Cell viability should be greater than 90%.
+
+1. Warm all reagents to 37°C prior to use.
+
+2. Remove growth medium from cells.
+
+3. Thoroughly rinse cell monolayers with 5 mL Ca²⁺- and Mg²⁺-free PBS per T75 flask or 100 mm dish. Gently rock the flask (or dish), allowing the solution to bathe the cells for 30 to 60 seconds at room temperature. Aspirate the rinse solution and discard.
+
+4. Repeat step 3.
+
+5. Add approximately 5 mL of Cell Dissociation Buffer per T75 flask or 100 mm dish and gently bathe cells by rocking at room temperature for 1 to 2 minutes. You may check for dissociation under the microscope. Aspirate solution and discard.
+
+6. Firmly tap flask or dish against palm of hand to dislodge cells. If cells do not detach quickly, allow to sit at room temperature for another 2 to 5 minutes and tap flask against palm of hand again. This may be repeated for cells that are more strongly adherent (with 5 more milliliters of dissociation buffer). After the cells are visibly detached, add at least 5 mL of complete growth medium. Resuspend cells in growth medium.
+
+NON-ENZYMATIC DISSOCIATION REAGENTS
+================================================================================
+
+Catalog numbers: 13150016, 13151014, 15040066
+
+REFERENCES
+================================================================================
+
+Freshney, R. (1987) Culture of Animal Cells: A Manual of Basic Technique, p. 117, Alan R. Liss, Inc., New York.

biomni/tool/protocols/thermofisher/F-Actin Staining _ Thermo Fisher Scientific - US.txt

@@ -0,0 +1,43 @@
+PROTOCOL: F-Actin Staining
+SOURCE: https://www.thermofisher.com/us/en/home/life-science/cell-analysis/cell-analysis-learning-center/molecular-probes-school-of-fluorescence/imaging-basics/protocols-troubleshooting/protocols/f-actin-staining.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+This protocol provides general instructions for labeling F-actin in fixed and permeabilized cells. Actin is one of the most abundant proteins found in cells and can be labeled with a fluorophore very easily when the monomers form a certain type of microfilament, referred to as F-actin. When phalloidin, a molecule that binds F-actin, is conjugated to a fluorophore, it becomes a very useful tool for cell biologists who are using fluorescence microscopy to study their cells. Labeling F-actin can help show the overall shape and structure of the cell and provide context for other fluorescent labels.
+
+Protocol tip: If you're using fluorescent phalloidin to visualize actin in your experiment, you'll get better results if you perm for 20 minutes instead of 15.
+
+MATERIALS REQUIRED
+================================================================================
+
+• Cells that have been fixed and permeabilized (See Fix, Perm, and Block protocol)
+• Phosphate-buffered saline (PBS)
+• Phalloidin conjugated to fluorophore
+• Fluorescence microscope with filter set matched to your fluorophore
+
+PROTOCOL
+================================================================================
+
+F-actin dyes—basic labeling protocol
+
+For fixed and permeabilized cells. Can be used before or after immunolabeling.
+
+The volumes given in this protocol are good for a single well in a 6-well vessel or a single 35 mm vessel.
+
+1. Prepare 1 mL F-actin staining solution in PBS from conjugated fluorophore. Phalloidins are typically very water soluble and stain F-actin at 100–200 nM concentrations. If you are optimizing for dye concentration, you will need to prepare a staining solution for each concentration you want to test.
+
+2. Remove PBS from fixed and permeabilized cells.
+
+3. Add 1 mL of staining solution.
+
+4. Incubate for 20 minutes at room temperature.
+
+5. Remove dye solution.
+
+6. Wash 3 times with PBS.
+
+7. Image cells.
+
+8. Optional: To preserve your sample for long-term storage, you can also mount your sample in mounting medium. Before you start your staining experiment, it's a good idea to check that your fluorophores are compatible with the mounting medium you want to use.

biomni/tool/protocols/thermofisher/Fluorescence In Situ Hybridization using TSA™ _ Thermo Fisher Scientific - US.txt

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+PROTOCOL: Fluorescence In Situ Hybridization using TSA™
+SOURCE: Thermo Fisher Scientific - US
+URL: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/microscopy-protocol/fluorescent-in-situ-hybridization-using-tsa.html
+================================================================================
+
+Introduction
+This protocol describes steps for fluorescent in situ hybridization (FISH) to Drosophila embryos using Tyramide Signal Amplification (TSA™), and was developed in collaboration with Ethan Bier and Dave Kosman at UCSD. This protocol is demonstrated with Drosophila embryos, but should be easily amenable to any specimen or tissue. TSA provides enzymatic fluorescent signal amplification that is ideal for low-abundance DNA and RNA targets. This approach utilizes DNA or RNA probes that have been labeled with a hapten such as biotin, fluorescein, digoxigenin, or DNP. In brief, the haptenylated probe is hybridized to the specimen and then TSA performed to detect the hybridized probe in situ. The hapten is detected in one of two possible approaches. The hapten can be detected by first applying an appropriate anti-hapten primary antibody followed by the horse radish peroxidase (HRP)-secondary antibody provided in the kit. The Alexa Fluor dye tyramide is then incubated with the specimen to detect the hybridized probe. Alternatively, haptens can be developed with anti-hapten–antibody-HRP or streptavidin-HRP conjugates. For example, biotinylated probes are detected with a streptavidin HRP conjugate provided in the kit. Similarly, anti-fluorescein–HRP conjugates can be used to detect fluorescein-labeled probes (but not included in the kit). Our TSA kits are available with a selection of dye tyramides and HRP conjugates; please consult our full product line to determine which best fits your experimental design. Importantly, TSA can be performed sequentially for multiple haptens by simply inactivating one HRP conjugate before applying another. This approach is ideal for multiplex FISH studies.
+
+Before You Begin
+Preparing Solutions for FISH in Drosophila Embryos
+
+PBT (2 L)
+- 200 mL 10X PBS
+- 2 mL Tween 20
+- To 2 L with water
+
+PBT+5% formaldehyde (15 mL)—make fresh
+- 12.86 mL PBT
+- 2.14 mL 37% formaldehyde
+
+Proteinase K (10 mg/mL stock)
+- 100 µL 20 mg/mL proteinase K stock
+- 100 µL water
+- Divide into 10 µl aliquots and store at ≤–20°C.
+
+2X carbonate buffer for fragmentation (10 mL)
+- 127.2 mg sodium carbonate
+- 67.21 mg sodium bicarbonate
+- To 10 mL with water and pH to 10.2 with NaOH; divide into 100 µL aliquots and store at ≤–20°C.
+
+Stop Solution: 0.2 M Sodium acetate, pH 6.0 (10 mL)
+- 164.06 mg sodium acetate
+- To 10 mL with water and pH to 6.0 with acetic acid; divide into 100 µL aliquots and store at ≤–20°C.
+
+Heparin (10 mg/mL stock in PCR-grade water)
+- Divide into 250 µL aliquots and store at ≤–20°C.
+
+Hybridization solution (150 mL)
+- 75 mL formamide
+- 37.5 mL 20X SSC
+- 1.5 mL herring sperm DNA (10 mg/mL)
+- 750 µL heparin (10 mg/mL)
+- 150 µL Tween 20
+- 35.1 mL water
+- Store at 4°C.
+
+Preparing Solutions for TSA™ Procedure
+Labeled Tyramide Solution
+Add 150 µL of high-quality anhydrous DMSO to one vial labeled tyramide (the vial comes from the Molecular Probes™ Tyramide Signal Amplification Kit). Divide into 10 µL aliquots; store at ≤–20°C until required.
+
+Procedure
+1. Fixing the Specimens
+Note: All steps in this section are to be performed at room temperature unless stated otherwise. Prior to aspirating and discarding solutions from the tube containing the embryos, ensure that the embryos are settled to the bottom to avoid discarding them.
+
+1.1. Place embryos in a 1.5 mL tube and bring to 1 mL with 100% ethanol (EtOH)
+1.2. Rock for 5 minutes.
+1.3. Pour off most of the EtOH (leave ~100 µL).
+1.4. Add ~900 µL of xylenes (so that the final solution is 90% xylenes and 10% EtOH).
+1.5. Rock the tube for 1–2 hours.
+1.6. Aspirate and discard the solution.
+1.7. Wash the embryos 2 times with EtOH (1 mL each wash).
+1.8. Aspirate and discard the last EtOH wash.
+1.9. Add 1 mL fresh EtOH and rock for 5 minutes.
+1.10. Aspirate and discard the EtOH.
+1.11. Wash the embryos 2 times with 100% methanol (MeOH), 1 mL fresh MeOH each wash.
+1.12. Add 1 mL fresh MeOH and rock for 5 minutes.
+1.13. Aspirate and discard the MeOH.
+1.14. Add 500 µL MeOH and 500 µL of PBT+5% formaldehyde to the embryos and rock for 5 minutes.
+1.15. Wash the embryos one time with 1 mL PBT+5% formaldehyde.
+1.16. Aspirate and discard the PBT+5% formaldehyde.
+1.17. Add 1 mL fresh PBT+5% formaldehyde and rock for 25 minutes.
+1.18. Aspirate and discard the PBT+5% formaldehyde.
+1.19. Add 1 mL PBT and rock for 5 minutes.
+1.20. Repeat the PBT wash step 3 more times (including 5 minutes' rocking each time) using fresh PBT each time.
+1.21. Remove an aliquot of 10 mg/mL stock solution proteinase K from the freezer and dilute 1:1000 in PBT.
+1.22. Aspirate and discard the last PBS wash from the embryos, add 1 mL of the 10 µg/mL proteinase K solution, and rock for 5–7 minutes.
+1.23. Aspirate and discard the proteinase K solution.
+1.24. Wash the embryos two times with PBT.
+1.25. Remove and discard the last PBT wash, add 1 mL fresh PBT, and rock for 5 minutes.
+1.26. Remove and discard the PBT.
+1.27. Add 1 mL of PBT+5% formaldehyde and rock for 25 minutes.
+1.28. Aspirate and discard the PBT+5% formaldehyde.
+1.29. Add 1 mL PBT and rock for 5 minutes.
+1.30. Repeat the PBT wash step 3 more times (including 5 minutes' rocking each time) using fresh PBT each time.
+1.31. Make a solution of 50% PBT/50% hybridization solution (preparation above).
+1.32. Aspirate and discard the last PBT wash, add 1 mL of 50% PBT/50% hybridization solution, and rock for 10 minutes.
+1.33. Aspirate and discard the 50% PBT/50% hybridization solution.
+1.34. Add 1 mL hybridization solution and incubate in a 55°C water bath for at least 1 hour (up to 3 hours).
+1.35. If hybridization will be performed the same day, keep the embryos at 55°C until ready for use. Otherwise store the embryos at ≤–20°C until you are ready for the hybridization procedure, but note that extended storage in hybridization solution (more than two weeks) results in degraded morphology over time.
+
+2. Optional: Fragmenting the Probe
+This optional step is intended to reduce the length of the RNA probe to 200–300 bases in order to increase penetration and improve S/N. This step may not be required for all probes, but should be considered as a means to increase S/N. As the incubation duration will determine the final RNA length, a few different times should be tested and the RNA product analyzed by gel electrophoresis.
+
+2.1. Add 25 µL of 2X carbonate buffer (preparation for 2X carbonate buffer above) to 10 µL of probe.
+2.2. Mix well and incubate at 42°C for 30 minutes.
+2.3. To the fragmented probe add:
+   - 50 µL of stop solution (preparation for stop solution above)
+   - 10 µL of 3 M sodium acetate, pH 5.2
+   - 1 µL of glycogen
+   - 300 µL ice cold 100% EtOH
+2.4. Mix well and store at ≤–20°C for 30 minutes (or longer).
+2.5. Pellet the nucleic acid by centrifugation.
+2.6. Wash the pellet with 400 µL of 70% ice cold ethanol.
+2.7. Air dry to remove residual ethanol, but do not allow the pellet to dry out completely.
+2.8. Add 5 µL of water and resuspend the pellet.
+2.9. Measure the concentration of a small amount of the resuspended probe (≤2 µL).
+2.10. Add hybridization buffer to the rest of the probe such that it is at a final concentration somewhere in the range of 5–50 ng/µL.
+2.11. Store the probe stock at ≤–20°C until you are ready for the hybridization procedure.
+
+3. Hybridizing the Probe
+3.1. Place the tube containing the embryos in a 55°C water bath until equilibrated.
+3.2. Aliquot the embryos into individual 1.5 mL tubes (~25 µL of stirred-up embryos per tube). Make as many tubes as you have RNA probes for.
+3.3. For each probe, make a solution containing 100 ng of probe in 100 µL of hybridization solution in a 1.5 mL tube.
+3.4. Incubate the tube(s) containing the probe at 65°C for 10 minutes to denature the probe.
+3.5. Aspirate and discard most of the hybridization solution from the tube(s) containing the embryos.
+3.6. Add 100 µL of denatured probe to the tube containing the embryos, and stir the embryos by tapping the tube.
+3.7. Incubate for 20–24 hours at 55°C.
+Note: it is recommended that the embryos be stirred once (by tapping the tube) ~10 minutes into the 55°C incubation period and once again before leaving them overnight. Ensure that the water bath is covered during the incubation to minimize condensation in the tube.
+
+4. Post-hybridization
+4.1. Stir up the embryos by tapping the tube with your finger.
+4.2. Aspirate and discard as much of the hybridization solution as possible.
+4.3. Add 1 mL of fresh hybridization solution, equilibrated to 55°C, to the embryos and gently invert tube 3-4 times.
+4.4. Place tube in a 55°C water bath and incubate for 5 minutes.
+4.5. Aspirate and discard the hybridization solution.
+4.6. Add 1 mL of fresh hybridization solution, equilibrated to 55°C, to the embryos and incubate at 55°C for 30 minutes. During this incubation, periodically remove the tube from the bath and tap it to stir up the embryos.
+4.7. Aspirate and discard the hybridization solution (ensure that the embryos have settled before aspiration).
+4.8. Add 1 mL of fresh hybridization solution, equilibrated to 55°C, to the embryos and incubate at 55°C for 30 minutes. During this incubation, periodically remove the tube from the bath and tap it to stir up the embryos.
+4.9. Make a solution of 50% PBT/50% hybridization solution (preparation above).
+Note: Steps 4.10 through 4.15 are performed at room temperature.
+4.10. Aspirate and discard the hybridization solution, add 1 mL of 50% PBT/50% hybridization solution, and rock for 10 minutes.
+4.11. Aspirate and discard the 50% PBT/50% hybridization solution.
+4.12. Add 1 mL PBT.
+4.13. Aspirate and discard the PBT.
+4.14. Repeat the PBT wash step 3 more times with 5 minutes' rocking each time using fresh PBT each time.
+
+5. TSA™ Procedure
+All steps in the TSA Procedure are performed at room temperature.
+5.1. Just before you begin the TSA procedure:
+   - Aspirate and discard the last PBT wash (from step 4.14).
+   - Add 1 mL of freshly prepared PBT+blocking reagent and rock for 30–60 minutes at room temperature or 37°C.
+5.2. Aspirate and discard the PBT+blocking reagent.
+5.3. Add antibody (1 mg/mL) diluted 1:100 in PBT+blocking reagent and rock for 2 hours at room temperature, protected from light. For example, add 4 µL antibody into 396 µL of PBT+blocking reagent. The final concentration should be 1 µg/mL.
+5.4. Aspirate and discard the antibody solution.
+5.5. Add 1 mL PBT.
+5.6. Aspirate and discard the PBT.
+5.7. Wash with PBT 4 times with 15 minutes' rocking each time, protected from light using fresh PBT each time.
+5.8. Aspirate and discard the PBT.
+5.9. Make mixture of 300 µL amplification buffer working solution (from step 5.1) and 3 µL of labeled tyramide solution (from freezer stock) and add it to the embryos.
+5.10. Rock for 10–15 minutes.
+5.11. Aspirate and discard the tyramide solution.
+5.12. Wash the embryos 3 times with PBT (1 mL each wash).
+5.13. Aspirate and discard the last PBT wash.
+5.14. Add 1 mL PBT and rock for 5 minutes at room temperature.
+5.15. If you are ready to image, aspirate and discard the PBT, add 1 mL 70% glycerol/30% PBT and rock for 10 minutes. Store at ≤–20°C until ready to image. We recommend that the embryos are stored no longer than 3 weeks prior to imaging.
+5.16. Aspirate and discard the 70% glycerol/30% PBT.
+5.17. Add 20 µL ProLong Gold antifade reagent (Invitrogen Cat. no. S36931 or S36935) to the embryos and deposit the sample onto a slide.
+5.18. Cover with coverslip and image using appropriate filters (visit probes.thermofisher.com for the spectral characteristics of the dye you are using).
+
+Reference
+1. Science 305, 846 (2004).

biomni/tool/protocols/thermofisher/Functional Mitochondrial Staining _ Thermo Fisher Scientific - US.txt

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+PROTOCOL: Functional Mitochondrial Staining
+SOURCE: https://www.thermofisher.com/us/en/home/life-science/cell-analysis/cell-analysis-learning-center/molecular-probes-school-of-fluorescence/imaging-basics/protocols-troubleshooting/protocols/functional-mitochondrial-staining.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+This protocol provides basic instructions to perform a fluorescence-based assay to detect functional mitochondria. Healthy mitochondrial membranes maintain a difference in electrical potential between the interior and exterior of the organelle, referred to as a membrane potential. Tetramethylrhodamine, methyl ester (TMRM) is a cell-permeant dye that accumulates in active mitochondria with intact membrane potentials. If the cells are healthy and have functioning mitochondria, the signal will be bright. Upon loss of the mitochondrial membrane potential, TMRM accumulation will cease and the signal will dim or disappear.
+
+Reminder: This assay is for live cells.
+
+MATERIALS REQUIRED
+================================================================================
+
+• Live cells
+• Complete medium (whatever you are growing your cells in)
+• Tetramethylrhodamine, methyl ester (TMRM)
+• Phosphate-buffered saline (PBS), or any other saline-based buffer
+• TRITC filter set for your fluorescence microscope
+
+PROTOCOL
+================================================================================
+
+Functional mitochondrial staining—basic labeling protocol
+
+Protocol tip: TMRM is often supplied as a powder. It's useful to make a stock solution in DMSO and store it at –20°C (for example, add 5 mL DMSO to 25 mg of TMRM to make a 10 mM stock solution). Once you're ready to use your stock, because TMRM is used at a low working concentration, you will need to make intermediate dilutions.
+
+• To make 50 µM intermediate dilution of TMRM → 1 µL 10 mM TMRM + 200 µL complete medium
+• To make 250 nM staining solution of TMRM → 5 µL 50 µM TMRM + 1 mL complete medium
+
+The volumes given in this protocol are good for a single well in a 6-well vessel or a single 35 mm vessel.
+
+1. Prepare 1 mL of 250 nM TMRM staining solution in complete medium. If you are optimizing for dye concentration, you will need to prepare a staining solution for each concentration you want to test.
+
+2. Remove media from live cells.
+
+3. Add the TMRM staining solution.
+
+4. Incubate for 30 minutes at 37°C.
+
+5. Wash 3 times with PBS (or other clear buffer).
+
+6. Image using TRITC filter.

biomni/tool/protocols/thermofisher/Hoechst 33342 Protocol for Imaging _ Thermo Fisher Scientific - US.txt

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+PROTOCOL: Hoechst 33342 Protocol for Imaging
+SOURCE: Thermo Fisher Scientific - US
+URL: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/protocols/hoechst-33342-protocol-imaging.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+Hoechst 33342 is a cell-permeant blue-fluorescent DNA stain that binds to the minor groove of double-stranded DNA, with preference for A-T rich regions. Unlike DAPI, Hoechst 33342 can readily enter live cells, making it useful for both live and fixed cell imaging applications. It is commonly used for nuclear staining and cell counting in fluorescence microscopy.
+
+APPLICATIONS
+================================================================================
+
+This protocol can be used for:
+• Nuclear staining in live or fixed cells for fluorescence microscopy
+• Cell counting and nuclear morphology analysis
+• Counterstaining in multicolor immunofluorescence experiments
+• Live cell imaging of nuclear structure
+
+This protocol should not be used for:
+• Flow cytometry (use Hoechst 33342 Protocol for HCA Instruments instead)
+
+MATERIALS REQUIRED
+================================================================================
+
+• Cells (live or fixed, on coverslips, slides, or in multiwell plates)
+• Hoechst 33342 stain (e.g., Hoechst 33342, trihydrochloride, trihydrate, Cat. No. H3570)
+• Phosphate-buffered saline (PBS) or cell culture medium
+• Mounting medium (for fixed cells, e.g., ProLong Gold Antifade Mountant)
+• Fluorescence microscope with DAPI/Hoechst filter set (excitation ~350 nm, emission ~461 nm)
+
+PROTOCOL
+================================================================================
+
+For live cells
+
+1. Prepare Hoechst 33342 stock solution:
+   - Dissolve Hoechst 33342 powder in water or DMSO to make a 1-10 mg/mL stock solution.
+   - Store at 2-8°C, protected from light. Stable for at least 6 months.
+
+2. Prepare working solution:
+   - Dilute stock to 1-10 µg/mL in cell culture medium or PBS.
+   - For most applications, 1-5 µg/mL is sufficient.
+
+3. Remove cell culture medium from cells.
+
+4. Add Hoechst 33342 working solution to cover cells.
+
+5. Incubate for 10-30 minutes at 37°C (or room temperature), protected from light.
+
+6. For live cell imaging:
+   - Remove staining solution and replace with fresh culture medium or imaging buffer.
+   - Image immediately using a fluorescence microscope.
+
+7. For fixed cell imaging:
+   - After staining, fix cells according to your standard protocol.
+   - Wash 2-3 times with PBS.
+   - Mount and image.
+
+For fixed cells
+
+1. Fix and permeabilize cells according to your standard protocol.
+
+2. Wash cells 2-3 times with PBS.
+
+3. Prepare Hoechst 33342 working solution (1-5 µg/mL in PBS).
+
+4. Remove PBS from cells.
+
+5. Add Hoechst 33342 working solution to cover cells.
+
+6. Incubate for 10-20 minutes at room temperature, protected from light.
+
+7. Wash 2-3 times with PBS.
+
+8. Mount cells in mounting medium (without DAPI if using Hoechst 33342).
+
+9. Image cells using a fluorescence microscope with DAPI/Hoechst filter set.
+
+SPECTRAL INFORMATION
+================================================================================
+
+Hoechst 33342:
+• Excitation maximum: ~350 nm (UV)
+• Emission maximum: ~461 nm (blue)
+• Standard filter set: DAPI/Hoechst
+• Compatible with: FITC, TRITC, Texas Red, Alexa Fluor dyes
+
+PROTOCOL TIPS
+================================================================================
+
+• Hoechst 33342 is light-sensitive; protect from light during staining and storage
+• Optimal concentration may vary by cell type; start with 1 µg/mL and adjust as needed
+• For live cells, use lower concentrations (1-2 µg/mL) to minimize toxicity
+• Higher concentrations may be needed for some tissue sections or thick samples
+• Hoechst 33342 can be used in combination with other fluorescent stains
+• For long-term live cell imaging, minimize exposure time and use lower concentrations
+• Fixed cells stained with Hoechst 33342 can be stored at 2-8°C, protected from light, for several weeks

biomni/tool/protocols/thermofisher/Immunolabeling Using Secondary Antibodies _ Thermo Fisher Scientific - US.txt

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+PROTOCOL: Immunolabeling Using Secondary Antibodies
+SOURCE: https://www.thermofisher.com/us/en/home/life-science/cell-analysis/cell-analysis-learning-center/molecular-probes-school-of-fluorescence/imaging-basics/protocols-troubleshooting/protocols/immunolabeling-using-secondary-antibodies.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+This protocol provides general instructions for immunolabeling cells with a primary and secondary antibody. Immunolabeling is a technique for fluorescently labeling a specific biological target within a sample using an antibody. You'll also see immunolabeling referred to as both ICC (immunocytochemistry) and IHC (immunohistochemistry), and sometimes even antibody labeling.
+
+MATERIALS REQUIRED
+================================================================================
+
+• Fixed, permeabilized, and blocked cells (See Fix, Perm, and Block protocol)
+• Primary antibody
+• Secondary antibody conjugated with fluorophore of choice
+• Phosphate-buffered saline (PBS)
+• Antibody staining solution.
+
+Note: If there is a recommendation from the manufacturer, then use the solution recommended to dilute your antibodies. Usually the antibody staining solution is made with diluted blocking solution (1% blocking solution in PBS) or just PBS.
+
+• Fluorescence microscope with filter set matched to your fluorophore
+
+PREPARING YOUR PRIMARY ANTIBODY STAINING SOLUTION
+================================================================================
+
+Before you begin, you will need to prepare your primary antibody staining solution. The antibody you have purchased will usually come with recommendations for its use, but if it doesn't, a good place to start is at 1 μg/mL, which is usually a 1:1,000 dilution from your tube if the primary antibody concentration is 1 mg/mL. That means for 1 mL of staining solution, you would add 1 μL of antibody to 1 mL of PBS or blocking solution.
+
+A dilution series can help you identify optimal signal:background conditions
+
+If you're using an antibody for the first time, it never hurts to try a few different concentrations to find the one that gives you the highest signal and lowest background for your target. Try a simple series of dilutions (1:100, 1:250, 1:500, 1:750, and 1:1,000) for your primary antibody while holding the secondary antibody concentration constant. You can do this experiment in a 96-well plate in order to quickly find the optimal concentration (see vessel considerations).
+
+Most secondary antibodies are used between 1 and 10 μg/mL. A good starting concentration for a typical secondary antibody in that concentration range would be a dilution of 1:1,000. If you find your staining to be extremely bright, or that you have too much background, you can always try a higher dilution (from 1:2,000 to 1:10,000).
+
+Use controls to make sure your signal is not an artifact
+
+If possible, it's a good idea to have both a positive and negative control for an experiment that uses immunolabeling. A positive control would be an alternate sample, in addition to your experimental sample, that contains high levels of your target. A positive control will verify that you have done the immunolabeling properly. A negative control would be one where the sample is incubated only with secondary antibody (omitting the primary antibody step); this will show you the level of background fluorescence (nonspecific fluorescent signal) that is coming from your secondary antibody.
+
+PROTOCOL
+================================================================================
+
+General immunofluorescence protocol using secondary detection
+
+1. Remove the blocking solution from your sample.
+
+2. Add enough primary antibody staining solution to cover your samples, except for your negative control, which you will incubate without primary antibody, but with the same buffer you diluted your antibodies in.
+
+3. Incubate for 1 hr at room temperature or overnight at 4°C.
+
+Note: Once you have completed this step, you can put your experiment in the refrigerator until the following day.
+
+4. Remove the primary antibody staining solution from your samples.
+
+5. Wash 3 times with PBS.
+
+6. Add enough secondary antibody staining solution to cover your samples, including your negative control.
+
+7. Incubate for 30 min to 1 hr at room temperature.
+
+8. Wash 3 times with PBS.
+
+9. Optional: Go to fluorescent labeling if you wish to stain your cells with other markers and haven't done it yet.
+
+10. Image cells.

biomni/tool/protocols/thermofisher/Mitochondrial Membrane Potential Assay Protocol _ Thermo Fisher Scientific - US.txt

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+PROTOCOL: Mitochondrial Membrane Potential Assay Protocol
+SOURCE: Thermo Fisher Scientific - US
+URL: https://www.thermofisher.com/us/en/home/references/protocols/cell-and-tissue-analysis/apoptosis-protocol.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+Mitochondrial membrane potential (ΔΨm) is a key indicator of mitochondrial health and cellular viability. Loss of mitochondrial membrane potential is an early event in apoptosis. This protocol describes methods for measuring mitochondrial membrane potential using fluorescent dyes such as JC-1, TMRM, or MitoTracker dyes.
+
+APPLICATIONS
+================================================================================
+
+This protocol can be used for:
+• Detecting early apoptosis
+• Assessing mitochondrial health
+• Screening for mitochondrial toxins
+• Studying mitochondrial function
+
+MATERIALS REQUIRED
+================================================================================
+
+• Cells in culture
+• JC-1 dye, TMRM, or MitoTracker dye
+• Cell culture medium
+• Phosphate-buffered saline (PBS)
+• Fluorescence microscope or flow cytometer
+• Optional: CCCP (carbonyl cyanide m-chlorophenyl hydrazone) as positive control
+
+PROTOCOL
+================================================================================
+
+Using JC-1 dye (ratiometric indicator)
+
+JC-1 exists as a monomer (green fluorescence) at low membrane potential and forms aggregates (red fluorescence) at high membrane potential.
+
+1. Prepare cells in appropriate culture vessel.
+
+2. Prepare JC-1 working solution:
+   - For JC-1 powder: Dissolve in DMSO to make 1-5 mg/mL stock. Store at -20°C.
+   - Working solution: Dilute to 1-10 µg/mL in cell culture medium or PBS.
+
+3. Remove cell culture medium.
+
+4. Add JC-1 working solution to cells.
+
+5. Incubate for 15-30 minutes at 37°C, protected from light.
+
+6. Wash cells 2-3 times with PBS or culture medium.
+
+7. For live cell imaging:
+   - Replace with fresh culture medium or imaging buffer.
+   - Image immediately using fluorescence microscope.
+
+8. For fixed cells:
+   - Fix cells according to standard protocol after JC-1 staining.
+   - Mount and image.
+
+9. Image analysis:
+   - Green fluorescence (monomer): excitation ~485 nm, emission ~530 nm
+   - Red fluorescence (aggregate): excitation ~535 nm, emission ~590 nm
+   - Calculate red/green ratio: higher ratio indicates higher membrane potential
+
+Using TMRM (Tetramethylrhodamine, methyl ester)
+
+1. Prepare TMRM stock solution (1-10 mM in DMSO).
+
+2. Prepare working solution (50-250 nM in cell culture medium).
+
+3. Add TMRM working solution to cells.
+
+4. Incubate for 20-30 minutes at 37°C, protected from light.
+
+5. Wash cells 2-3 times with PBS.
+
+6. Image using TRITC filter set (excitation ~540 nm, emission ~565 nm).
+
+7. Higher fluorescence intensity indicates higher membrane potential.
+
+PROTOCOL TIPS
+================================================================================
+
+• Include positive control (cells treated with CCCP or other mitochondrial uncoupler) to confirm assay is working
+• Optimize dye concentration and incubation time for your cell type
+• Protect from light during staining and imaging
+• For JC-1, use ratiometric analysis (red/green) for more accurate results
+• Some dyes may be toxic to cells at high concentrations; use lowest effective concentration
+• For flow cytometry, use appropriate controls and compensation settings
+• Fixed cells can be used, but live cell imaging provides more dynamic information

biomni/tool/protocols/thermofisher/Mounting Coverslips _ Thermo Fisher Scientific - US.txt

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+PROTOCOL: Mounting Coverslips
+SOURCE: https://www.thermofisher.com/us/en/home/life-science/cell-analysis/cell-analysis-learning-center/molecular-probes-school-of-fluorescence/imaging-basics/protocols-troubleshooting/protocols/mounting-coverslips.html
+================================================================================
+
+OVERVIEW
+================================================================================
+
+Formulations of mounting media that can add favorable properties such as optimizing the refractive index to match that of glass, preventing photobleaching, or preserving samples for long-term storage are widely available. Keep in mind that hard-setting mountants require time to "cure" or harden.
+
+MATERIALS REQUIRED
+================================================================================
+
+• Fixed, permeabilized, blocked, and stained sample on coverslip in a saline-based buffer solution such as PBS
+• Glass microscope slide
+• Mounting medium
+
+PROTOCOL
+================================================================================
+
+Try to limit the formation of bubbles in your mounting medium. Some bubbles are inevitable, and you can image around them as long as there are only a few.
+
+To avoid bubbles:
+• Don't shake or invert your bottle of mounting medium.
+• Clear bubbles from the tip of your applicator (i.e., from the tip of the pipette or dropper bottle) by squeezing a little bit of mounting medium onto a lab tissue before applying mounting medium to your slide
+
+Clean the surface of the non-sample side of the coverslip prior to imaging. Fingerprints, residual salts, and other contaminants can diminish image quality.
+
+1. Apply a small amount of mounting medium to the surface of the slide; try to use an amount that will just fill the space under the coverslip.
+
+2. Remove coverslip containing the sample from the buffer.
+
+3. Blot excess buffer or solvent from the non-sample surface of the coverslip (or allow it to air-dry and then remove the salt residue before imaging).
+
+4. Slowly tip the coverslip onto the mounting medium, and avoid creating bubbles as you lower it into place.
+
+5. Follow manufacturer's directions for curing time.
+
+6. Optional: Seal the coverslip if desired.
+
+7. Image.