Team:NU Kazakhstan/Protocols

Valet — A free HTML5 Template by FREEHTML5.CO

Chlamydomonas reinhardtii protocols


  1. Resuspend and take 100μL of algae from the old solution of TAP+arg
  2. Place into fresh 50 mL TAP+arg
  3. Check OD at 750nm


  1. Put 15 g of Agar (SIGMA-ALDRICH) in 2 L Erlenmeyer flask and fill up with 1.5 L of dH2O
  2. Put in magnetic stirrer rod and place on magnetic plate
  3. Wait for 3 hours
  4. Take Buchner funnel and place a filter paper on it
  5. Attach it to vacuum filtering flask, turn on vacuum pump and seal filter paper on the surface of Buchner funnel (pour a little bit of dH2O)
  6. After vigorous swirling pour about 250 mL of agar with water in the Buchner funnel
  7. After several minutes, agar will dry a little bit on filter paper
  8. Take it out from the funnel very accurately and place on the dish
  9. Discard all liquid from vacuum filtering flask into the rack
  10. Repeat steps 6-9 until all agar is filtered
  11. Carefully take all dried agar from filter papers back into washed 2 L Erlenmeyer flask
  12. Fill it with dH2O again
  13. Repeat 7 washes with dH2O, 1 wash with 10% bleach, 1 wash with 70% ethanol and 1 final wash with dH2O. Total 10 washes
  14. After final wash, collect all agar from filter papers and place it onto some glass or plastic flat surface and leave it to dry for 2 days


  1. Weight 29.4185 g of K2Cr2O7 and put into 500 mL glass bottle
  2. Firstly put 150 mL of pure TAP and dissolve solid particles
  3. Then perform pH adjustment using 10 M NaOH solution
  4. Filter solution using large vacuum filters into sterile autoclaved 500 mL glass bottle


  1. Put all reagents in accordance to following recipe:
    [Gorman, D.S., and R.P. Levine (1965) Proc. Natl. Acad. Sci. USA 54, 1665-1669]
    Beware!!! FINAL PH SHOULD BE 7.0. Adjust if necessary.
  2. Phosphate Buffer II
  3. Autoclave. Let it cool down
  4. Filter
  5. Add arginine (final concentration should be 10 ug/mL)
  6. Adjust pH:
    1. 400 mL of TAP medium need to be prepared for autoclaving with amount of salts identical for 500 mL. Add such amount of salts, so that it will reach proper concentration after adjusting total volume to 500 mL
    2. The same with arginine. Final concentration of arginine need to be 50 mg/L
    3. Adjust pH to 7.0-7.3 using pH meter and titrating it with HCl
    4. Arginine is added, pH is adjusted, now finalize volume up to 500 mL


  1. Take 1.5 g of washed agar and fill up with 100 mL of TAP medium (depending on what type of plates you want: with or without chromium, arginine, acetate) in 500 mL glass bottle
  2. Place into large autoclave
  3. Pour agar into 6-8 60 mm petri dishes


  1. Loopful of culture from each tube and inoculate it in 100mL in an Erlenmeyer flask of liquid media
  2. Add arginine for CW- strains and antibiotics upon necessary
  3. Leave it under continuous illumination and shaking on the shaker at room temperature in the hood
  4. Close the top with sterile cotton and aluminum foil


  1. When cells reach OD 0.5-0.7, harvest them by centrifugation at 2,500 rpm for 5 minutes. Discard the supernatant and carefully remove all liquid as much as possible. Note: Cells must be in early log phase and harvested gently. If the cell concentration is <1 × 106 cells/mL, you can still harvest the cells without significantly affecting the transformation efficiency. If the cell concentration exceeds 3 × 106 cells/mL, discard the cells and start a new culture
  2. Resuspend the cell pellet in 10 mL of GeneArtTM MAX EfficiencyTM Transformation Reagent and centrifuge at 2,500 rpm for 5 minutes. Discard the supernatant and carefully remove all liquid as much as possible
  3. Resuspend the cell pellet again in 10 mL of GeneArtTM MAX EfficiencyTM Transformation Reagent, and centrifuge the cells once more at 2,500 rpm for 5 minutes
  4. Resuspend the cell pellet in 1 mL GeneArtTM MAX EfficiencyTM Transformation Reagent
  5. Add 1 μg of linearized DNA per 100 μL of cell suspension and incubate at 2°C–8°C for 5 minutes
  6. Fill the NeonTM Tube with 3 mL of ice-cold E2 buffer and insert it into the NeonTM Pipette Station until you hear a click. Note: After 2–3 shocks, E2 buffer needs to be chilled on ice again
  7. Set electroporation parameters on the NeonTM device as follows: Voltage: 2300V, Pulse Width: 3ms, Pulse Number: 3
  8. Pipette up 100 μL of the DNA-cell mix in the 100-μL NeonTM Tip and insert the tip into the NeonTM Tube in the pipette station until you hear a click
  9. Press Start on the touchscreen to deliver the electric pulse
  10. Eject the electroporated cells into a 15-mL centrifuge tube (chilled on ice) and allow the cells to recover on the bench for 15 minutes. Add 4 mL of TAP-40 mM sucrose solution at room temperature to the cells and incubate them in the algal chamber overnight
  11. The next day, centrifuge the cells at 2,500 rpm for 5 minutes, discard 3.8 mL of the supernatant, and resuspend the cells in the remaining 200 μL of TAP-40 mM sucrose solution
  12. Spread 200 μL of the cell suspension on a TAP-agar-ZeocinTM plate using disposable cell spreaders or glass plating beads to spread the cells evenly. Make sure the plates do not have condensation on them
  13. Place the plates agar side down in the algal growth chamber set to 26°C and 50 μE m–2 s–1. Do not stack the plates to ensure continuous and even illumination


  1. Take 250 ul of culture, centrifuge at max speed for 10 min at 4 degrees
  2. Quickly remove supernatant and make sure it does not contain parts of the cells pellet
  3. Resuspend cell pellet in 20 ul of 0.2% Triton
  4. Heat the tubes with cells at 98C for 10 min
  5. Centrifuge at 14 000 g and take up the supernatant
  6. Extract the supernatant with hexane and remove lower aqueous layer into separate tubes. This layer contains DNA. Make sure you do not take up hexane
  7. Set up PCR reaction using 2ul of DNA
  8. Use standard reaction protocol for Q5 polymerase and Phusion Master Mix with primers ChrR pchlamy

PCR machine settings :


General/E.Coli protocols


  1. Add 37g of nutrient agar to 400 mL of the distilled water
  2. Autoclave


  1. Suspend 25 grams in 1000 ml distilled water
  2. Heat if necessary to dissolve the medium completely
  3. Sterilize by autoclaving at 15 lbs pressure (121°C) for 15 minutes
  4. Dispense as desired


  1. Beforehand prepare 0.1M CaCl2 and 0.1M CaCl2 + 15% glycerol solutions
  2. Take 1 colony of DH5-alpha strain of E.coli from LB plates and inoculate in 10mL of LB in 50 ml Falcon tube
  3. Place the tube with DH5-alpha E.coli strains in LB into incubator at 37 oC, until OD600 reaches 0.2-0.3, shaking at 250 rpm
  4. Once the OD600 reaches 0.2-0.3, place the tube with DH5-alpha strain on ice for 15 min. Keep in ice solutions of 0.1M CaCl2 and 0.1M CaCl2 + 15% glycerol too
  5. Centrifuge cells at 4 oC for 10 min
  6. Resuspend pellet with 3 ml of 0.1M CaCl2 and put on ice for 30 min
  7. Centrifuge cells again and resuspend in 300 µL of 0.1M CaCl2 + 15% glycerol
  8. Prepare 50 µL aliquots of resulting solution in separate Eppendorf tubes
  9. Place aliquots in Cold room at -80 oC


was carried out according to IGEM Protocols

  1. Resuspend DNA in selected wells in the Distribution Kit with 10µl dH20. Pipet up and down several times, let sit for a few minutes. Resuspension will be red from cresol red dye
  2. Label 1.5ml tubes with part name or well location. Fill lab ice bucket with ice, and pre-chill 1.5ml tubes (one tube for each transformation, including your control) in a floating foam tube rack
  3. Pipette 50µl of competent cells into 1.5ml tube: 50µl in a 1.5ml tube per transformation. Tubes should be labeled, pre-chilled, and in a floating tube rack for support. Keep all tubes on ice. Don’t forget a 1.5ml tube for your control
  4. Pipette 1µl of resuspended DNA into 1.5ml tube: Pipette from well into appropriately labeled tube. Gently pipette up and down a few times. Keep all tubes on ice
  5. Pipette 1µl of control DNA into 2ml tube: Pipette 1µl of 10pg/µl control into your control transformation. Gently pipette up and down a few times. Keep all tubes on ice
  6. Close 1.5ml tubes, incubate on ice for 30min: Tubes may be gently agitated/flicked to mix solution, but return to ice immediately
  7. Heat shock tubes at 42°C for 45 sec: 1.5ml tubes should be in a floating foam tube rack. Place in water bath to ensure the bottoms of the tubes are submerged. Timing is critical
  8. Incubate on ice for 5min: Return transformation tubes to ice bucket
  9. Pipette 950µl SOC media to each transformation: SOC should be stored at 4°C, but can be warmed to room temperature before use. Check for contamination
  10. Incubate at 37°C for 1 hours, shaking at 200-300rpm
  11. Pipette 100µL of each transformation onto petri plates Spread with sterilized spreader or glass beads immediately. This helps ensure that you will be able to pick out a single colony
  12. Spin down cells at 6800g for 3mins and discard 800µL of the supernatant. Resuspend the cells in the remaining 100µL, and pipette each transformation onto petri plates Spread with sterilized spreader or glass beads immediately. This increases the chance of getting colonies from lower concentration DNA samples
  13. Incubate transformations overnight (14-18hr) at 37°C: Incubate the plates upside down (agar side up). If incubated for too long, colonies may overgrow and the antibiotics may start to break down; un-transformed cells will begin to grow


was performed according to the Monarch® PCR & DNA Cleanup Kit (5 μg) Protocol (NEB #T1030)


was performed according to the Monarch® DNA Gel Extraction Kit | NEB


Isolate maxiprep plasmid DNA

  1. Equilibrate. Apply 30 mL Equilibration Buffer (EQ1) directly into the Filtration Cartridge, which is inserted into the Maxi Column. Allow the solution in the HiPure Filter Maxi Column to drain by gravity flow
  2. Harvest. Centrifuge the overnight LB culture at 4000xg for 10 minutes. Remove medium
  3. Resuspend. Add 10 ml Resuspension Buffer (R3) with RNase A to the cell pellet and gently shake the pellet until the cell suspension is homogenous
  4. Lyse. Add 10 ml Lysis Buffer (L7). Mix gently by inverting the capped tube until the mixture is homogenous. Do not vortex. Incubate the tube at room temperature for 5 minutes
  5. Precipitate. Add 10 ml Precipitation Buffer (N3). Mix immediately by inverting the tube until the mixture is homogenous. Do not vortex
  6. Clarify. Transfer the precipitated lysate into the column. Allow the lysate to filter through the column by gravity flow. Optional. Wash the column with 10 ml Wash Buffer (W8). Allow thebuffer to flow through the column by gravity flow
  7. Wash. Discard the inner filtration cartridge. Wash the column with 50 ml Wash buffer (W8). Discard the flow-through after the buffer drains
  8. Elute. Place a sterile 50-ml centrifuge tube under the HiPure Filter Column. Ass 15 ml Elution Buffer (E4) to the column. Allow the solution to drain by gravity flow. Discard the column. The elution tube contains the purified DNA

Precipitate using a centrifuge

  1. Precipitate. Add 10.5 ml isopropanol to the eluate. Mix well. Centrifuge the tube at >12,000xg for 50 minutes at 4C. Discard the supernatant
  2. Wash. Add 5ml 70% ethanol to the pellet. Centrifuge the tube at >12,000xg for 5 minutes at 4C. Remove the supernatant
  3. Resuspend. Air dry the pellet for 10 minutes. Add 500 ul (for high copy number plasmids) or 200 ul (for low copynumber plasmids) TE buffer to the ourified DNA. Store plasmid DNA at -20C


All centrifugation steps are performed at maximum speed (12,000–14,000 x g)

  1. Transfer an overnight culture (1–2 ml) of plasmid-containing cells to a microcentrifuge tube. Pellet the cells by centrifugation for 15–30 secs. Remove all of the supernatant by aspirating or pipeting
  2. Add 200 μl of the cell resuspension solution and vortex or pipet up and down until the cell pellet is completely resuspended
  3. Add 250 μl of the cell lysis solution and mix by gently inverting the capped tube about ten times (do not vortex). The solution should become viscous and slightly clear if cell lysis has occurred
  4. Add 250 μl of the neutralization solution and mix by gently inverting the capped tube about ten times (do not vortex). A visible precipitate should form
  5. Pellet the cell debris for 5 mins in a microcentrifuge. A compact white debris pellet will form along the side or at the bottom of the tube. The supernatant (cleared lysate) at this step contains the plasmid DNA
  6. While waiting for the centrifugation step at step 5, insert a spin filter into one of the 2 ml microcentrifuge wash tubes supplied with the kit. Mix the Quantum Prep matrix by vortexing or repeated shaking and inversion of the bottle to insure that it is completely suspended
  7. Transfer the cleared lysate (supernatant) from step 5 to a spin filter, add 200 μl of thoroughly suspended Quantum Prep matrix, then pipet up and down to mix. If you have multiple samples, transfer the lysates first, then add matrix and mix. When matrix has been added to all samples and mixed, centrifuge for 30 secs
  8. Remove the spin filter from the 2 ml tube, discard the filtrate at the bottom of the tube, and replace the spin filter in the same tube. Add 500 μl of wash buffer and wash the matrix by centrifugation for 30 seconds
  9. Remove the spin filter from the 2 ml tube, discard the filtrate at the bottom of the tube and replace the spin filter in the same tube. Add 500 μl of wash buffer and wash the matrix by centrifugation for a full 2 mins to remove residual traces of ethanol
  10. Remove the spin filter and discard the microcentrifuge tube. Place the spin filter in one of the 1.5 ml collection tubes supplied with the kit or in any standard 1.5 ml microcentrifuge tube that will accomodate the spin filter. Add 100 μl of deionized H2O or TE. Elute the DNA by centrifugation for 1 min at top speed
  11. Discard the spin filter and store the eluted DNA at -20°C


  1. Inoculate 50 ml of bacterial culture in appropriate selective media and incubate on shaker at 250 rpm at 37C overnight
  2. Next day transfer the bacterial culture into a 50 ml tube and spun at 4000xg at 4C for 10 minutes
  3. Discard the supernatant and resuspend the pellet in 2ml of resuspension buffer with 50ul/ml RNase (ThermoFisher #EN053) freshly added
  4. Add 2ml of lysis buffer to the bacterial suspension and invert the tube 3-4 times
  5. Incubate it at room temperature for 3 minutes
  6. Add 2ml of neutralization buffer and invert the tube 3-4 times
  7. Quickly distribute the bacterial lysate into 1.5ml centrifuge tubes (apprx. 4 tubes) by pouring, not pipetting
  8. Centrifuge at room temperature at 13,200xg for 10 minutes
  9. Collect supernatants in 15 ml tube and discard pellets
  10. Add 1x volume of 96% ethanol (apprx. 5ml) into the supernatant and mix it thoroughly for 5 seconds
  11. Load the sample-ethanol mix onto 5 spin-columns in three sequential (apprx.700ul) aliquots
  12. Spun the column for 30 seconds at 13,200xg after the addition of each aliquot
  13. After each spin discard the flow through and repeat the steps till the entire sample passed through the spin columns
  14. Wash the columns 2 times with 500ul of wash buffer
  15. After each wash spun them at 13,200xg at room temperature for 30 seconds
  16. Discard the flow through
  17. Centrifuge the empty columns one more time for 1.5 minutes to remove any residue buffer
  18. After this, discard old collection tube and put the column into a new tube
  19. Add 30-35ul of elution buffer to the column and incubate for 2 minutes
  20. Spun at 13,200xg at room temperature for 2 minutes
  21. Combine the eluted DNA from all columns in one tube (apprx. 175ul)
  22. After measuring the concentration store the samples at -20C


  1. Rinse Nanodrop with a ethanol and Kimtech paper towels
  2. Launch ND-8000 V2.2.1.
  3. Make sure that pedestals are clean
  4. Blank with 2ul of Nuclease Free water (work very accurately!)
  5. Blank with 2ul of buffer (Elution buffer after miniprep and gel extraction)
  6. Load 1ul of DNA (another person should mix the sample via pipetting up and down prior that: not vigorously!). Work very fast! Close the cover
  7. Choose wells and run
  8. Clean with ethanol and Kimtech paper towels after each use
  9. Place cover back on


  1. Prepare 1% agarose gel via adding 1g of agarose powder into 100ml of 1XTAE buffer
  2. Heat it till agarose is completely dissolved (Do not boil!)
  3. Add SYBR Safe as required (see the tube) in dark room
  4. Prepare the mould by wrapping it with aluminum foil
  5. Pour the solution into the mould and make sure that there are no bubbles
  6. Allow the solution to set (apprx. 15-20 minutes)
  7. Remove comb and load ladder and samples (5ul of DNA and 1 ul of Loading Dye for 6x dye)
  8. Run gel at 100V
  9. Visualise bands


was carried out according to NEB requirements.
For 50 uL reaction:

  1. Buffer (10x) : 5 uL (CutSmart)
  2. Enzyme 1: 0.5 uL
  3. Enzyme 2: 0.5 uL or dH2O
  4. DNA : 1000 /concentration (total 1000ng of DNA)
  5. ddH2O: fill up to 50 ul
  6. Put it in thermostat at 37 degrees for 1 hour, heat inactivate at 65/80 degrees for 20 min (ell enzymes we used required heat for inactivation, consult NEB website)


was carried out according to NEB protocol: “Ligation Protocol with T4 DNA Ligase (M0202)”

  1. Set up the reaction mixture in eppendorf tube (for 1:3 Vector-Insert): pro5
  2. Mix the reaction by pipetting up and down
  3. For sticky ends incubate it at 16C overnight
  4. Heat inactivate at 65C for 10 minutes
  5. Chill on ice and transform 1-5ul of reaction into 50ul of competent cells


Carried out in accordance to NEB “PCR Protocol for Taq DNA Polymerase with Standard Taq Buffer (M0273)”.


was conducted in accordance to Quan & Tian, 2011, “Circular polymerase extension cloning for high-throughput cloning of complex and combinatorial DNA libraries”

Colony PCR

Carried out by NEB protocols for Phusion Master Mix solution what we found as the best possible protocol for colony PCR. The general protocol is here.
However, we added some modifications in order to our protocol to increase the efficiency. Instead of using extracted DNA, we used a single colony, which was boiled for 10 min at 95C. Then, 1 uL of solution was used to set the general protocol. A single colony should be stirred rigorously and well (crucial point). In addition, we figured out the the addition of 3% DMSO is important to maximize the PCR.


  1. Revive 1 vial of frozen C. reinhardtii cells and inoclate 200-ml of TAP medium Culture the cells under standard consitions and keep monitoring their concentrations for 3 days
  2. When cell concentration reaches 106-2 x 106 cells/ml, harvest them by centrifugation at 2500 rpm. Discard the supernatant and carefully remove cell liquid as much as posible
  3. Resuspend the cell pellet in 10 ml of Gene Art Max Efficiency Transformation Reagent and centrifuge at 2500 rpm for 5 minutes. Discard the supernatant, and carefully remove all liquid
  4. Concentrate with Gene Art Max Efficiency Transformation Reagent to 108-3x108 cells/ml. Add 1 ml of the solution
  5. Add 1 ul of linearized DNA per 100ul of cell suspension and incubate at4 C for 5 minutes
  6. Fill the Neon Tube with 3 ml of ice-cold E2 buffer and insert it into the Neon Pipette station until you hear a click. Note: after 2-3 use, E2 buffer needs to bechilled at 4 C
  7. Ser electroporation parameter of the Neon Device as follows: Voltage 2300V, Pulse width 13 ms, Pulse Number 3
  8. Pipette up 100 ul of the DNA-cell mix in the 100 ml Neon Tip and insert the tip into Neon tube in the pippete station until you hear a click
  9. Press the Start button on the touch screen to deliver the electric pulse
  10. Eject the electroporated cells into a 15 ml centrifuge tube (chilled on ice) and allow the cells to recover on the bench for 15 minutes
  11. Add 4ml of TAP-40 mM sucrose solution at room remperature to the cells and incubate them in the algal chamber overnight
  12. The next day, centrifuge the cells at 2500 rpm for 5 minutes, discard 3.8 ml of the supernatant, and resuspend the cells in the remaining 20ul of TAP-40mM sucrose solution
  13. Spreas 200ul of the cell suspension on selective TAP-agar plates, and incubate in the algal chamber for 5-7 days


  1. First make the Phenol:Chloroform solution as follows. Mix Phenol:Chloroform:Isoamyl alcohol in 25:24:1 ratio
  2. Add one volume of cold Phenol:Chloroform vortex for a few minutes. Spin down for 10 minutes
  3. Take supernatant to a new microtube. First use a P1000, then a P200 to get as much as possible. Two clear phases should form. Pull up very slowly
  4. Now add one volume chloroform-phenol at room temperature, vortex for 1-2 min. and spin down (about 10 minutes)
  5. Pull off the aqueous layer again
  6. Add another volume of chloroform-phenol, vortex and then spin down. At this point there should be almost no protein left. Take supernatant slowly
  7. Now add chloroform (pure) one volume to the supernatant, vortex it and centrifuge for 10 min . Repeat the step twice
  8. Precipitate that, therefore take supernatant and pour to the new eppendorf tube. Add 1/10 volume of sodium acetate pH 5.2, followed by two volumes of ice-cold Ethanol and incubate at -20°C for overnight
  9. Now transfer the tube to -4°C, leave there for ten minutes and then spin down at 20,000 g for 10 minutes in a microcentrifuge
  10. Wash the pellet with cold 70% Ethanol. This means add some 70% Ethanol and shake then spin down
  11. Let the sample air-dry
  12. Resuspend in nuclease-free water
  13. Measure the concentration of the DNA at this point


Reagent setup

  • Fixer solution - dilute glacial acetic acid to 7.5% (v/v) with deionized water. Developer stop solution has the same recipe
  • Formaldehyde solution - prepare 15% (v/v) with ddH20. Store at room temperature
  • Silver solution - dissolve 0.1 g AgNO3 in 100 mL in ddH20. Always make a new fresh solution
  • Sodium thiosulfate stock solution - dissolve 0.2 g Na2S2O4 in 50 mL ddH20
  • Developer solution - dissolve 3.0 g Na2CO3 in 100 mL ddH20. Swirl it in the cold box. Store at 4°C
  • Developer stop solution - the same as fixer solution. Store at 4°C
  1. Nucleic acid fixation. Put the gel upside down and pour fixation solution so that it covers 5 mm the gel. Incubate on a rocker for 10 min
  2. Prepare fresh solutions. Add Na2S2O4 (50 ul) per 100 mL Developer solution
  3. Gel washing. Decant the solution and pour ddH20; rock for 2 minutes. Repeat the step 3 2 more times
  4. Formaldehyde pre-treatment. Add 5 mm of formaldehyde solution and incubate on a rocker for 10 min. Decant the solution
  5. Silver impregnation. Add sufficient solution (silver stain) to cover the gel. Incubate for 30 minutes on a rocker. Decant the silver. Wash the gel for 5 sec with ddH20
  6. Image development. Add the developer solution for 3 minutes. After, decant the solution
  7. Stopping the reaction. Add developer stop solution for 5-10 minutes Decant it. Wash with ddH20. The developer stop solution should be ice cold

The complete Western Blotting Protocol

Preparation of Total Cell Extracts by the AT Lysis Method

AT Buffer (final conc.)

  • Make up EDTA, EGTA, NaF, Na4P2O7, DTT as 10X stock, store at -20°C
  • Make up Hepes as 1 M stock, store at 4°C
  • Once made up, AT Buffer can be stored at -20°C indefinitely in small aliquots
  • Components marked * must be added freshly on day of use
  • To isolate cytosol only, try only 0.1% Triton X-100 and no glycerol

To prepare 100 mL of AT Buffer


To make extracts

  1. Aspirate media from 100 mm (60 mm) plate of cells and wash cells 2 times with ice-cold PBS or TD. After the second wash, add 1 mL of 1X PBS (or TD), scrape cells gently with a rubber scraper, and collect in microcentrifuge tube. Add another 0.5 mL of ice-cold PBS (or TD) to the plate, rescrape and pool with the first 1 mL scraping
  2. To collect cells, spin 3-4 min at about number 5 speed in microcentrifuge and aspirate as much of the PBS (or TD) as possible from the cell pellet when done
  3. Add 160 uL of ice-cold AT buffer (or 100 ul for 60 mm dish)
  4. Aspirate through a 27-gauge needle at least 5 times. Squirt liquid onto side of tube as you are aspirating up and down
  5. Add 5 uL of 5 M NaCl (to a final conc. of 150 mM NaCl) (or 3.1 uL of 5 M NaCl if working with cells scraped from 60 mm dish)
  6. Spin at top speed in microcentrifuge at 4°C for 30 min
  7. Transfer supernatant to a new tube. At this point, extracts can be stored at - 80°C

Preparation of Cell Extract by SDS/boiling Lysis

  1. Aspirate media from cells and wash cells 2 times with ice-cold TD buffer. Scrape off cells as above in about 1.5 mL of TD buffer (1 mL then 0.5 mL and then pool)
  2. To collect cells, spin for 5 min at approximately #5 speed in microcentrifuge and aspirate to remove as much TD as possible from the cell pellet
  3. Depending on the number of cells and the abundance of protein, add water, resuspend cells and then add an equal amount of 2X or 4X SDS sample buffer. For a 60 mm dish of cells, you may want to add about 100 uL of water and 100 uL of SDS sample buffer (and run about 10-20 uL on the gel after boiling)
  4. Mix sample well, and boil sample for 10 min at 95-100°C
  5. Extract is now ready for loading onto SDS gel or can be stored at -80°C (if stored frozen, reboil about 3 min before loading on SDS-polyacrylamide gel)

Protein Determination (OD 595)

  1. Dilute Bio-Rad solution to 1X (It comes as a 5X stock) e.g., 2 mL of Bio-Rad Protein Assay Dye Concentrate Reagent + 8 mL of distilled water
  2. Aliquot 1 mL of 1X Bio-Rad Protein Assay solution to microcentrifuge tubes. Of course, the number of centrifuge tubes depends on number of samples
  3. Set up the Bovine serum albumin (BSA) standard using the BSA stocks from New England Biolabs: 100X BSA stock is equal to 10 ug/uL, and 10X BSA stock contains 1 ug/uL. Therefore, to the aliquoted 1X Bio-Rad solution, add 0, 1, 2, 5, 10, 15, 20 ug by adding the following amounts of a given BSA stock to each tube: pro8
  4. For your samples, you want the Bio-Rad solution to turn blue within about a minute after addition of extract. Usually, this will happen if you add 2-5 uL of each cell extract to 1X Bio-Rad solution, and then mix by vortexing briefly and leave at room temperature for one minute
  5. When reaction has completed changing color, transfer 200 uL of reaction (i.e., either BSA standards or cell extracts) into a 96-well Falcon plate
  6. Read OD at 595 nm
  7. Find slope (ug/OD) of BSA standard, according to the following formula: Slope = ug (0+1+2+5+10+15+20)/absorbance (OD0+OD1+OD2+OD5+...+OD20)
  8. Multiply absorbance value of each cell extract sample by the calculated slope number (ug/OD). Divide that number by the volume of cell extract added to the 1X Bio-Rad solution (from step 4). Resulting value (ug/uL) is the protein concentration of cell extract. This will often be in the range of about 1-5 ug/uL
  9. Depending on the level of expression of the protein of interest you may need to load different amounts of cell extract. However, a good starting point is to use about 20 ug of cell extract on SDS gel. (To determine volume needed, divide 20 ug by the protein concentration of cell extract)


  1. Assemble the glass plate sandwich
  2. Prepare the resolving gel solution:

  • Pour the gel solution between the glass plates, leave about ¼ of the space free for the stacking gel
  • Carefully cover the top of the resolving gel with 50% isopropanol and wait until the resolving gel polymerizes (~30 min). A clear line will appear between the gel surface and the solution on top when polymerization is complete
  • Discard the isopropanol. Wash gently with double-distilled water
  • Prepare the stacking (upper gel): pro10
  • Pour the stacking gel solution
  • Insert combs. Allow the gel to polymerize for at least 60 min
  • Remove combs carefully. Put the gel into the electrophoresis tank, fill the tank (bottom and top reservoirs) with fresh 1X Tris-glycine-SDS Buffer, make sure that the gel wells are covered with the buffer
  • Load protein ladder/marker and probes
  • Set an appropriate voltage and current depending on how many gels you run
  • Increase the power when the dye front reaches the resolving gel
  • Stop the electrophoresis run when the dye front reaches the bottom of the gel. Disassemble the gel sandwich and proceed with Western Blot procedures

  • Transferring Proteins from SDS-Polyacrylamide Gels to Nitrocellulose

    10X Transfer Buffer
    [1X: 20 mM Tris, 0.15 M Glycine]
    24.22 g Tris
    112.6 g Glycine
    Add water to about 800 mL, once Tris and glycine are dissolved, adjust volume to 1000 mL.

    1X Transfer Buffer
    100 mL 10X Transfer Buffer
    200 mL Methanol
    700 mL Water
    1000 mL in total

    1X High protein Transfer Buffer
    100 mL 10X Transfer Buffer
    100 mL Methanol
    800 mL Water
    1000 mL in total
    *Wet Whatman paper and transfer apparatus sponges with 1X Transfer Buffer

    Prepare the blot sandwich as follows:

    1. Black side of transfer apparatus
    2. Two sheets of Whatman 3 MM paper
    3. Gel
    4. Nitrocellulose membrane (nitrocellulose also needs to wetted for about 30 sec in 1X Transfer Buffer before putting against gel). When laying nitrocellulose on gel, be sure to lay out smoothly and with no bubbles (use a glass dowel). Wear gloves and handle nitrocellulose with forceps
    5. Two sheets of Whatman 3 MM paper
    6. White side of transfer apparatus.
    7. Close apparatus tightly, and place in blotting apparatus. The nitrocellulose membrane should closest to the anode (red) part of the apparatus (for the cassette: Black to black - White to red).

    For minigels, use the mini-transfer apparatus, and transfer at 160 milliamps for 60 min (for higher molecular weight proteins, transfer for 2.5 hrs at 200 mA in cold room)

    Western Blotting with Pierce’s SuperSignal

    Solutions Required:

    • 25% Tween 20 [pour 10 mL of Tween 20 into a 50-mL falcon tube; add deionized water up to 40 mL]
    • 1X TBS
    • 1X TBS + 0.1% Tween
    • TMT: 1X TBST, 2% milk [100 mL: 5 g Carnation instant nonfat milk; TBST to 100 mL] Note: for some antisera, you will need to use 3% BSA-not milk-in the antibody dilution buffer)


    1. Blocking Step: Immerse nitrocellulose in TMT and shake slowly on platform for 1 hr at room temperature (or overnight at 4 degrees)
    2. Remove blocking solution and add diluted primary antibody. If the antibody is diluted in TBST-BSA, you will have to wash 3X in TBST for 5 min each before adding antibody.
    3. You should add enough primary antibody to just cover the gel. Very gently rotate the blot in primary antibody in PMT for 1 hr at room temperature. Make sure that during the shaking the liquid is continuously covering all of the blot: if you rotate it too quickly you can create a situation where the liquid is not circulating over the center of the blot --- this is bad. (Usually you can save the primary antibody after use and use it several times. In fact, it often gets better [cleaner] with extended use. Store the diluted antibody/milk solution at -20 degrees)
    4. Remove primary antibody. Wash the blot by shaking in an excess of TMT for 5 min. Repeat 4 times, i.e., 5X 5-min washes; in other words, these washing steps should take about 25-30 min
    5. Remove and discard the final wash solution, and add the secondary antibody. Slowly shake membrane in diluted secondary antibody for at least 1 hr. For goat anti-rabbit HRP, use 1:10,000-20,000 for Pierce reagent, i.e., 1uL in 10 mL or 20 mL of TMT. Make the dilution by first pouring the given volume of TMT (e.g., 10-20 mL) into a plastic tube, then add the secondary antibody, mix by inverting a few times and pour onto the nitrocellulose. Note, the dilution of the secondary varies for each primary antibody and the detection reagents that are used, and often must be determined experimentally
    6. Remove and discard the secondary antibody. Wash the blot in TBS-Tween for 4 washes for 10 min each
    7. Wash in 1X TBS for 2 washes for 5 min each
    8. Leave blot in TBS until ready to expose to substrate. To prepare for the addition of substrate, lift the nitrocellulose with forceps to drain excess TBS (but do not dry) and put onto Saran wrap with protein side up
    9. Prepare Pierce’s SuperSignal (Pierce SuperSignal WestDura Extended Substrate) just before you are ready to probe the blot. For each substrate, add an equal amount of reagent 1 and 2 to a 15 mL snap top tube and mix gently (for a small blot, use 0.5-1 mL of each reagent, i.e., 1-2 mL total should be sufficient to cover blot)
    10. Slowly pour the substrate solution onto the nitrocellulose membrane (protein side up). Leave on for 5 min (Pierce). While the substrate solution is on the membrane be sure to gently and carefully move the saran wrap so that the substrate continuously and fully moves across the membrane. To avoid any darkroom conflicts, it is usually a good idea to do the substrate addition in the darkroom, so that you will have ready access to the exposure and developing facilities. That is, take your blot upstairs in TBS, do the substrate addition in the darkroom, and then do your exposures
    11. After incubating with the substrate for 5 min, use forceps to lift blot, allow the excess substrate to drain, and place the blot onto a new piece of saran wrap. Place the protein side face down onto a smooth surface of saran wrap. Package the membrane by closing saran wrap on back
    12. Place the blot protein side up in a film cassette, close cassette tightly and expose to film. (Note: do not expose with white radiation enhancing screens in the cassette. These will cause fogging of the film especially on long Western blotting exposures.) The time of exposure will vary depending on the antibody, the abundance of the given protein, the amount of extract used, etc. To save time and film, it is advisable to perform multiple exposures on a single piece of film; this is done by moving the nitrocellulose membrane to a fresh part of the film and allowing to “expose” for varying lengths of time. One can usually get 6 exposures of a small filter per 8 x 10 inch piece of X-ray film. For example, to do 1, 5, 10, 30, 60, 120 sec exposures. You can cut the film in half if you only want to do 2-3 exposures, but do not cut the film smaller than this. If using a new antibody, and you do not know how long to expose the blot, a five minute exposure is usually sufficient to determine if you need to expose it longer or five minutes is too long. If you can see your proteins “glowing” in the dark room on the membrane, 1-10 second exposures should be sufficient.

    **The incubation times that are indicated above are the minimum times (i.e. you can leave the primary antibody incubation at 4 degrees overnight).
    It is advisable NOT to leave the secondary overnight, as that might increase the background

    1 M Hepes (pH 7.9)
    23.83 g Hepes
    Dissolve in 60 mL water, pH with 10 N NaOH to 7.9.
    Adjust volume to 100 mL
    1 M Na3VO4
    1.84 g sodium vanadate
    Dissolve in 7 mL water.
    Adjust volume to 10 mL.
    10X AT Stock
    500 uL DTT (from a 1 M stock)
    1 mL EDTA (from a 0.5 M stock)
    2 mL EGTA (from a 250 mM stock)
    0.420 g NaF
    4.46 g Na4P2O7 x 10 H2O
    Mix in about 80 mL of water and when mixed, adjust volume to 100 mL with water
    10X TBS
    80 g NaCl
    2 g KCl
    14.4 g Na2HPO4 - 7 H2O
    2.4 g KH2PO4
    Dissolve in 800 mL distilled water, then adjust volume to 1000 mL. Autoclave.
    1X TBS
    100 mL 10X TBS
    900 mL distilled water
    1X TMT with 5% milk
    5 g Carnation Instant Non-fat milk
    100 mL 1X TBS-Tween

    Stripping a Membrane and Reprobing with a Different Antibody

    1. Make 100 mL of stripping buffer
    2. Turn on the 55°C water bath and give it about 10-15 min to heat up
    3. Pour out old stripping buffer from stripping container
    4. Pour new stripping buffer into the container
    5. Put nitrocellulose into the stripping buffer, seal the container, and put it into the 55°C water bath with a red weight on top
    6. Incubate for 30 min
    7. Wash 3X in TBS-Tween for 5 min each
    8. Block with TMT for 1 hr at RT or overnight at 4°C
    9. Proceed to Western blotting with the different primary antibody
    10. When developing the blot, you may find that there is less signal on the blot, and so the best exposure may be significantly longer than expected. At times, 15-20 min of exposing to film is necessary to get a good signal on a reprobed Western blot.

    Chromium reduction assay protocol

    1. Transfer 25-mL of cultures of algae from the flask into the 50-mL tubes
    2. Wash cultures 3 times with PBS (pH7). In total 4 tubes
    3. Right after washing step, adjust volume to 4-mL and let it stand on ice for 20 minutes
    4. While the samples on ice, start sonication step. Use the following options: amplitude 25%, frequency, 120 Watt, 25kHz, 30 seconds work, 30 seconds rest, total time 4 minutes
    5. Immediately after sonication add protease inhibitor solution HALT
    6. Centrifuge for 3 minutes at 4700xg
    7. Filter all samples using 0.22um filters
    8. Adjust all samples’ pH to 5
    9. Using the nanodrop find the protein concentration in each filtered sample
    10. Assemble reaction mixture:
      • Filtered sample with 0.4 mg of protein content
      • 100mM Cr(VI) in NaOAc – 1uL
      • 10mM NADH – 10uL
      • NaOAc – fill to reach total volume of 1mL
      • Temperature conditions - 35 degree Celsius
    11. From each reaction mixture take 97.8uL of mixture, then add 872uL of acetate, 20uL of DPC solution, 10uL HCl and pour all of these into the cuvette. Measure OD 540
    12. Perform 11th step each 15 minutes during 1 hour