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Eisosomes
We applied out methods and tools to investigate the composition and the dynamics of Eisosomes. We tagged two proteins Pil1 and Lsp1 with the our modular tags and, purified and identified several proteins. Using reciprocal tagging we confirmed that some of the identified proteins co-localize with Pil1 and Lsp1, which indicates that they could be essential components of Eisosomes.
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APC
We purified yeast APC and mapped phosphorylation sites on several subunits: Apc1, Cdc16, Cdc23, Cdc26 and Cdc 27 using a modular mass spectrometric tool. We now puriffy cell-cycle specific APC complexes to profile the composition and the abundances of the phosphorylation sites on the subunits during the different stages of the cell cycle.
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Nuclear periphery proteins.
We study protein composition and cell-cycle dynamics of a number of nuclear periphery proteins from yeast and human cells. We use cell-cycle sub-cellular localization and protein-protein interaction data as clues to uncover the processes, in which these proteins are involved.
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Discovery and characterization of new substrates of Separase.
We about to initiate a project with the aim to discover and characterize the new substrates of separase. The technique and tools we develop could be extremely helpful for achieving this goal.
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Use prOTOF mass spectrometer to measure the tryptic map spectra.
version 20.12.06
1. The prOTOF mass spectrometer should indicate that it is ready for the analysis by a green light, “TOF Vacuum”, on the front panel.
2. Wake up the prOTOF computer by pressing the computer switch button quickly. Login and type a password.
3. Open TOFWORKS. Login as a tofworksuser and type a password.
4. Remove a target cassete from the instrument target chamber. Remove the blank metal MALDI target from the cassette and insert your magnetic target deposited on the thin metal plate (you should find it and leave it in the table drawer) . Be sure to close the camber completely by pressing the top hook all the way down.
5. Define your target either as new or as already existing one. This operation starts the sequence of the target insertion into the instrument. Wait until ready (~ 3-5 min), all for light are green (Instrument status-green, TOF vacuum - green, Plate Loaded - green, Plate Chamber Vacuum - green).
6.Switch on the small tv. In few moments you should see your target on the screen of the tv. Wait until the target is inserted completely into the instrument.
7. File/check plate alignment -> calibrate the plate alignment by marking two dots, X 2/ OK
8. Use the default instrument method to measure the spectra.
9. Give a name for the sample of the current sample spot.
10. Measure the spectrum of a 100 femtomole of a calibration peptide mixture. Acquire (scan the laser spot)/add to sum/ sum + current. Save the spectrum by going to the next layer and pressing “pick peaks”. Peaking peaks deposits the spectrum in the spectra data base.
11. Calibrate the instrument by measuring the spectrum of a peptide mixture. Use m/z values of ions at 1046.542 and 2456.199.
Acquire (scan the laser spot)/add to sum/ sum + current/ pick peaks/ calibrate:
Calibrant – calibrate. Set peaks found at m/z 1046.xxx to correct m/z 1046.5423, and peak found at m/z 2465.xxx to the correct m/z 2465.199. Usually the differences between the old values and the new ones are just few millimasses. Apply to all samples.
12. Go to Acquire data/Acquire (scan laser spot over your sample)/add to sum/ sum + current/ pick peaks/
13. Spectra are saved in the computer database after “pick peaking” procedure.
14. Extract the the collected spectra with the program “prOTOF extractor”. Give the spectra the appropriate names (suggestion: keep the sample position in the file name, example: B01_your file name.txt)
NOTE: Do not connect prOTOF computer to the internet! Use a flash memory to transfer your data.
Spectra processing
1. Use your computer or a nearby computer for analysis of your data. Login as a user and type a password.
Note: this computer is connected to the internet.
2. Use “m/z” program to find and label all ion peaks of interest. Suggestion: use spectrum smoothing procedure (use coefficient 3) and then the spectrum enhancement procedure. Use “A” - automatic mode of peak picking. Check manually whether all peaks are picked properly, and monoisotopic M+H peaks are selected. Add or remove peaks as necessary.
3. Copy the list of peaks in clipboard and then in the “Notepad” program. Save as a text file. Keep the name of the file short, representative of an analyzed spot. Do not use spaces in the file name.
4. These list of peaks may be used for ID of proteins based on the “tryptic mapping”.
Use Xproteo search engine or some other engine of your choice. This pre-screen helps to evaluate the quality of your sample, initial ID of proteins and, in general, the status of your project at this point.
Use MALDI-LTQ mass spectrometer to measure MS/MS spectra on all peaks detected in the tryptic maps.
1 . Copy the files containing the list of masses onto a flash drive. Transfer the data files to the computer of the MALDI-LTQ mass spectrometer.
2. Use AutoMSMS program to generate the script for the LTQ mass spectrometer to perform MS/MS analysis on all peaks from the given data file.
7. Chose the data file(s).
8. Define: num of laser shots per spot 5, if you decide to run the LTQ in completely automatic mode. Or use num of laser shots per spot 100, if you would like to have a manual control over the position of the laser spot on your sample. Leave other paramers as default. Run the program on all selected files. This program will create method files for analysis of each samples. Each method files “knows” what peaks to analyse in each sample and “how” to do it (what is a peak selection widht, activation energy and time etc).
9. Insert your magnetic MALDI target onto the plate holder and insert into of the LTQ mass spectrometer.
10. After all methods are generated run them either one-by one, in a semi-automatic mode usine Tune program. Or use Xcalibur program to define the sequence of analysis of all selected sample spots.
11. After the analysis is complete, generate the “DTA” files from data.raw files. Use the DTA converter program. (Select M0.3 and S0 tags for conversion). Transfer you data to the analysis computer. You can search and identify you proteins based on the accurate m/z of the parent peptides and their MS/MS spectra using several data bases. Use Xproteo, Mascot or some other search engines to ID the proteins.
NOTE: Do not connect vMALDI-LTQ computer to the internet! Use a flash memory to transfer your data.
We use double affinity tags, 3xFLAG-6xH or 3xMyc-8xH, for purification of proteins and protein complexes (see our IP protocol). The use of 3xFLAG or 3xMyc tags are usually sufficient to obtain highly enriched protein complexes. However, an additional tandem purification step can be very useful to remove an excess of the elution peptide and concentrate the complexes on the metal chelating resin. After the final wash of the chelating resin, the purified proteins can be efficiently eluted for separating by SDS-PAGE or left on the beads for on bead digestion.
Here is the protocol for digestion of proteins and protein complexes on the metal chelating resin.
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1.Wash the cobalt (or some other metal) chalating beads two times with 1 ml of 50mM ammonium bicarbonate buffer. The purpose of this step is to remove the detergent used in the IP buffer as completely as possible.
2. Add 5-10 ul of 1pmol/ul trypsin solution in 50 mM ammonium bicarbonate to the beads. Incubate 5-30min; collect the supernatant and incubate it further for 5-12 hours. Limited digestion may help to minimize elution of nonspecifically bound proteins (this still remains to be proven). Otherwise, add trypsin and digest on the beads all the time (6-12 hours)
3. Analyze 1/3 or ½ of the digest (3-5 ul) directly by a MALDI modular MS tool or HPLC-ESI.
Note: Filter or spin down the digest before loading a portion into an HPLC column. This step is necessary to minimize the chances of blocking the HPLC tubings with small micropaticles of metal chelating resin.
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Version 25.5.07
All operations, at least before stopping digestion, are preferably done in a clean dust free environment (air clean workstation, clean room etc.).
1. Cut the gel band of interest. Do not cut a sharp thin pieces. The width of the gel piece should be ~1 mm or more.
Note: There is a common notion of cutting a gel piece as close to its staining borders, which is not helpful or even wrong. The resolving power of all common gels is not high. Proteins usually migrate as rather wide bands. Gel staining however, frequently reveals a “tip of an iceberg”, and not the real spacial distribution of a protein in the gel.
When dealing with entire gel lane, slice a gel lane onto ~10-40 1-, 2- or 3-mm pieces with a Mickle gel slicer or any suitable cutter. On a copy of a gel picture mark the beginning, the end, and some intermediate positions of the slicer blade for future cross-reference and a gel slicing calibration. Use tweezers with a thin flat tips to pick up the pieces and to chop them onto ~1×1x1 mm pieces. All manipulation with gel pieces can be easily performed using a clean white or black plate. After slicing and dicing the gel pieces, transfer them directly into the 1.6 ml tubes, which were labeled and pre-filled beforehand with a ~1.5 ml of a distaining solution. For example, for coomassie stain use 1/1 v/v of 50mM ammonium bicarbonate and MeOH.
2. Distain for necessary period of time in ~1.5 ml distain solution with agitation. For example, for 10-20 min with Zn-distain, if gel was stained with Zn-stain, or for 30min-3 hours in coommasie distain solution (freshly made 50 mM ammonim bicarbonate in 50% MeOH, if the gel was stained with a colloidal coommasie. All reagents are the highest purity available). After distaining, remove solution as completely as possible.
Optional: Reduce and alkylate the proteins in the gel if needed at this point. Wash well the gel pieces after completion of the alkylation reaction.
3. Dehydrate the bands well. For this add 1ml of acetonitrile to all gel pieces and slightly agitate the vial for 10-15 min. All gel pieces should become very white color. Remove acetonitrile.
4. Dry the gel pieces in by leaving the tubes open. Optional: microwave for 1 min and let it dry.
Note: When the gel pieces become very dry, they have tendency to pop out of the tubes under the influence of static electricity that might be generated on the tubes after drying.
5. Dissolve well an aliquot of trypsin (2.5 ug) in 100 -200 uL of 50mM ammonium in a freshly made and filtered (0.45um filter) bicarbonate buffer, pH ~8. Add 5-10 ul of trypsin solution to gel pieces depending on an estimated volume of initial gel piece. Add just enough of trypsin solution to keep a gel piece slightly “thirsty” upon rehydration. Put the tubes on ice for 30-45 min ( to prevent self digestion of trypsin while gel rehydration)
6. After incubation on ice is finished, add ~10-20 ul of the same digestion buffer, but without trypsin.
7. Digest for 3-12 hours at 37 C.
8. Stop digestion and extract peptides by addition of 30-40 uL 7% formic acid/ 0.1% TFA in water. Also add ~1-2 ul of Porous 50 beads from a stock kept in methanol (beads/MeOH = ~1/5, v/v). The hydrophobic beads will act like a pump absorbing all peptides that diffuse out of the gel pieces.
Note: Amount of beads added should later create ~100-400 nL volume of bead column in the tip of a Gelloader tip or a Zip tip.
Gently shake the pieces of gel in the presence of beads for 3-12 hours in a medium speed shaker. For longer extraction times, ~ 4-12 h, should be done at 4C, to minimize hydrolysis of -D-P- bonds.
9. After extraction step, collect the beads in the bigger gel loader pipette trying to avoid any small gel pieces. By doing so, press the tip against the bottom of the vial and shake from side to side. While collecting aspirate several times to elute any beads that could stack to the pieces of gel or the walls of the vial. Transfer the collected beds into the Gelloader pipette the tip with a frit made from C8 membrane (3M). Collect the beads by gentle pushing the liquid and create a C18 column at the end of the tip. When processing many gel pieces use a centriguge designed to accommodate 30 GelLoader pipette columns on the flat rotor.
10. Wash the column with 5-20 uL of 0.1% TFA.
11. Elute with matrix solution of DHB dissolved in 60% methanol, 1-2% acetic acid.
If using 4hcca as a matrix, elute peptides with a solution with high content of organic solvent (60-70%+0.1% TFA or 2% acetic acid) directly to the target and then add the 4hcca matrix.
12. In case of using 4hcca as matrix, wash the final spot 2 times with 5-7 ul drops of 10% MeOH/0.1% TFA solution. Leave a droplet for 10-30 sec and remove the liquid by suction from a vacuum line.
1. Grow several liters of cells overnight to the density 2-5×10e7 cells/ml
2. Spin down cells in 1 liter bottles at 5000rpm 15-20’ 4ºC
3. Remove supernatant and collect cells with up to 50ml of ice cold water into 50ml conical tube
4. Spin down cells for 1’ at 4000rcf and remove the supernatant.
5. Add 0.5-1ml of IP buffer(see the recipe below) containing protease inhibitor cocktail. Use a spatula to re-suspend
6. Slowly pour the cells into a new 50 ml tube filled with liquid nitrogen and emerged in it. The slower the procedure, the smaller size of the frozen pellets, and lesser chance of possible sudden boiling of liquid nitrogen.
7. Important: Make several holes in a cap with a razor blade to exhaust nitrogen. This step is necessary to avoid explosion of the tubes because of the high pressure of the nitrogen vapors.
8. Pour off excess liquid nitrogen through the holes (use protective glasses and insulating gloves or forceps). Cells can be maintained at –80ºC for a long time at this stage.
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9. Put pellets in the screw-top jars previously immersed in the liquid nitrogen until it stops boiling. Insert the jars into a Retsch Mixer Mill MM301, and beat for 5 times, 3’ at 30 beats per second. Between each beating immerse the jars in liquid nitrogen to cool. Efficiency of cell breakage can be observed in a microscope.
10. Transfer a cell powder back into 50 ml tube pre-cooled in liquid nitrogen. Be sure that there is no residual liquid nitrogen left in the tube. Cell powder can be maintained at –80ºC for long time at this stage.
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11. For 1 IP experiment: Re-suspend 1-5 g of broken cells in 5-10ml IP buffer +0.1% Tween 20 +Cocktail of protease inhibitors + kinase inhibitors.
12. Let the tube thaw on ice for 10-15 min and then use a tissue homogenizer to mix. Mix quickly and cool before runs.
13. Solubilization step: incubate rotating 45’-60’ @ 4ºC
14. Spin down at full speed for 1’-5’ @ 4ºC
15. Transfer sup into new 10ml tube/s
16. Add 5-10 mg of Dynal beads with immobilized antibody to each sample.
17. Rotate 15-30’ 4ºC
18. Collect the beads using big magnet. Discard all sup except for 1ml – transfer it to a new 2ml eppendorf tube.
19. Wash 3X with IP buffer. Change to a new 1.65 ml tube.
20. Add 200µl 3XFLAG (from stock of 200µg/ml in 50mM Tris-HCl pH=7.4, 150mM NaCl))
21. Elute rotating for 15-30’ @ 4ºC
22. Transfer 200µl of supernatant to new 1.65 ml tube.
23. Add 1000µl IP buffer + 20µl Cobalt Talon beads from a commercial stock.
24. Rotate 4ºC 15’
25. Remove sup
26. Wash 2X1ml IP buffer, change the tube. Use few seconds spin to remove residual amount of washing liquid.
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Separate sample onto two parts by separating the liquid containing the Talon beads.
For separating the purified proteins by SDS-PAGE process as follows:
NOTE – FROM NOW ON BE VERY CAREFUL TO AVOID CONTAMINATING YOUR SAMPLE WITH KERATINS
27. Add 10µl Sample buffer containing 200mM Imidazole without reducing agent! Its presence rusts the Co beads.
28. Incubate 15’ 37ºC rotating
29. collect the sup using magnets
30. Add reducing agent and reduce for 15-30’ at 60 C.
31. Run SDS-PAGE use Micro-Gel 15’ or any suitable gel electrophoresis.
FOR ON BEADS DIGESTION
27a. Wash the remainig Co-Talon beads two times with 1 ml of 50mM ammonium bicarbonate buffer. And change the tube.
28a. Add 10 ul of 1pmol’ul trypsin solution in 50 mM ambic to the beads. Incubate 5-30 min; collect the sup and incubate further for 5-12 hours. Limited digestion may help to minimize elution of nonspecifically bound proteins (?, remains to be proven) Another option is just add trypsin and digest on the beads all time (6-12 hours)
29. Analyze 1/3 or ½ of the digest directly by MALDI or HPLC-ESI.
30. For HPLC-ESI, spin down the sample using high speed centrifuge to minimize the chance of blocking the tubes of an HPLC system.
We develop and build new mass spectrometric tools to advance our research. One recent development is a A Novel High‐Capacity Ion Trap‐Quadrupole Tandem Mass Spectrometer constructed in collaboration between our laboratory and Brian Chait’s laboratory at the Rockefeller University. This mass spectrometer was built as a prove of the principle for increasing the efficiency of linked scan analysis by 100-1000 fold.
Based on this prototype instrument, we currently construct and build a fully functional model of such a tandem mass spectrometer here at the UCSF. When build, this new tool will be indispensable for or detecting modification sites on the proteins and profiling their abundances with unprecedented speed and sensitivity.
When we study yeast, we use a homologous recombination to incorporate our modular tag at the C-terminus of the protein of interest (see for example Knop M et al Yeast 1999, 15, 963-972) at the C-terminus of the genomic sequence of the protein. The tagged proteins are expressed under the control of endogenous promoters.
To express the proteins of interests in human cells we use an in-house modified Flp-In in vivo recombination system from Invitrogen. This system allows us to produce stable cell lines expressing the tagged version of the proteins under the control of a tetracycline inducible promoter. Both cell systems are useful for observing protein localization and purifying protein complexes for their detailed characterization. In the future, however, we would like to add more cell types expressing of the studied proteins.
We designed and tested a modular epitope system for genomic tagging of the proteins. Our modular tag contains one of the versions of a fluorescent protein for in vivo localization and a small double affinity tag for immunopurification.
A modular approach allows us to decoupled the two types of experiments. We do not rely on the availability of a good antibody against a fluorescent protein for affinity purification. Instead we focus on optimizing the performance of each module for studying the dynamics of protein localization and interactions.
Here is a set of vectors we developed for tagging the proteins with a modular tag. The tag contains one of the versions of a monomeric fluorescent protein (GFP (S65T), EYFP, mOrange, mCitrine, Cherry etc.) fused at the C-terminus to a small double affinity tag for immunopurification (3xFLAG-6xH, 3xMyc-8xH, 4Strep-8xH, etc). The sequences of the vectors can be found here.
