Staining Protocol and Tips for Submitting Sypro Ruby Stained Gels for Protein Identification

Staining Protocol and Tips for Submitting Sypro Ruby
                         Stained Gels for Protein Identification

We will not accept any samples without first discussing the details of your sample. Please call or set up a
meeting to discuss their sample. Contact Melissa Sondej at masondej@ucla.edu or 310-206-0432.

Tips and Information

A. Avoiding Keratin Contamination
   The environment we work in is highly polluted with keratin that came from our heads, skin, nails,
   fingerprints, woolen clothing, and etc. Other rich sources of keratin contamination include paper and
   cardboard containers. Since mass spectrometers detect proteins at femtomolar levels, the slightest
   contamination of keratin will result in keratin being identified as the most abundant protein versus being
   able to identify the proteins that really interest you. A key factor in avoiding keratin contamination is to
   avoid contaminating your sample with dust which adheres to keratin. Therefore, use a scrupulously
   clean work surface and wear a clean lab coat to perform as much of the steps as possible from sample
   preparation all the way to the in-gel trypsin digestion. Next, keep in mind where your gloves have
   touched. Your gloves should only touch your sample. We cannot tell you the numbers of times we have
   seen people touch areas that are highly contaminated with keratin such as drawer handles, computer
   mouses and keyboards, and even their face/glasses and then touch their samples/gels. If you find it
   hard to take your gloves on and off, layer a second pair to touch dirty things. Also, make sure you keep
   the lids closed as much as possible on tubes/plates, tip boxes and solutions in order to avoid dusting
   from falling in/on them.

B. Other Types of Contamination
   We have seen other types of protein contamination such as bovine serum albumin and milk proteins.
   These usually come from a staining tray which was previously used to develop a western. Therefore we
   ask you to use a brand new staining tray. These contaminations can also come from reagent bottles
   and beaker. Therefore, be careful that you don’t make up your Coomassie blue stain, fixing and
   destaining solutions, and etc. in glassware that for instance has been used to make blocking solution
   for a western blot. If you don’t know the history of your glassware and you cannot use disposable
   plasticware, then soak your glassware overnight in Micro-90 detergent (VWR ) to remove any
   contaminating proteins. Make sure you rinse your glassware extremely well or the residual detergent
   can interfere with the mass spectrometry analysis.

C. Gloves
   You must use powder-free nitrile gloves. Never use latex gloves since they are composed of natural
   rubber which contains significant amounts of keratin and other proteinaceous materials. Also never use
   gloves containing aloe since this is a proteinaceous extract.

D. HEPA-filtered laminar-airflow hood
   If you are having severe keratin contamination even after doing all the things listed above, you may
   want to perform as much of your gel work as possible underneath a HEPA-filtered laminar-airflow hood.
   This work includes opening your gels, changing the fixer, staining and destaining solutions, and cutting
   your bands underneath a HEPA-filtered laminar-airflow hood. Before starting your experiment, wipe
   down the hood and equipment with ethanol to remove any keratin contamination. Working in a hood is
   not absolutely required but since gels are sticky and act like keratin magnets, it provides the best dust-
   free environment.

E. Pre-Cast Gels
   We highly advised you to purchase pre-cast gels. The gels you typically cast in your own lab are highly
   contaminated with keratin. If you are going to cast your own gel, you will need to prepare your gel 18-24
   hrs in advance to allow maximal polymerization. Also you will need to clean your glass plates with the
   following cleaning steps to remove protein contamination such as keratin.
        1) Soak the plates overnight in 5% Decon Contrad 70 liquid detergent (Thermo Fisher Scientific
           catalog number 04-355).
        2) Scrub the plates with a Kimwipe, and then rinse the plates thoroughly with nanopure equivalent
           water.
        3) Soak the plates for 1 hr in 1% HCl.
        4) Scrub the plates with a Kimwipe, and then rinse the plates thoroughly with nanopure equivalent
           water.
        5) Dry the plates with a Kimwipe.
        6) Store the plates in a clean plastic bag or container until use.

F. Molecular Weight Standards
   Sypro Ruby cannot stain Pre-stained Protein Molecular Weight standards. Therefore, make sure you
   use an unstained standard when running your gel.

G. Gel Electrophoresis
   For protein identification, you need to get rid of any band smearing since it cross contaminates your
   bands. One way to increase the sharpness of your bands, is to decrease your voltage down to 100 V
   for the electrophoresis run. This decreases the amount of heat that is generated during the run. Also,
   you may want to run your gel at 40 V for the first 10 min to allow better stacking of the proteins in the
   stacking gel.

H. Staining Trays
   Inivtrogen recommends using either polypropylene (i.e. a pipet tip box) or polycarbonate containers.
   These high-density plastics adsorb minimal amounts of the dye (examples include Servin’ Saver®
   and Stain Shield® containers from Rubbermaid). For the best results, use containers dedicated for
   SYPRO Ruby gel staining to minimize dye cross-contamination or other artifacts. Glass dishes are not
   recommended. [This section comes from the Invitrogen Sypro Ruby Mannual.]

I.   Cleaning Staining Trays
     Staining containers should be meticulously clean.
         1) Scrub the tray with soap and a kimwipe.
         2) Rinse the tray thoroughly with distilled water. Make sure you rinse away all the soap or it will
            interfere with the mass spec analysis.
         3) Scrub the tray with ethanol and a kimwipe.
         4) Rinse the tray thoroughly with distilled water.

J. Multiplex Staining
   This technology allows you to sequentially stain one gel with first a phosphosprotein stain (i.e.
   Invitrogen Pro-Q Diamond or Perkin-Elmer Phos-tag stain), followed by a glycoprotein stain (i.e.
   Invitrogen Pro-Q Emerald), and finally with a total protein stain (i.e. Invitrogen Sypro Ruby). The gel is
   imaged after each of the staining methods. Between stains, rinse the gel 2X 5 min in MilliQ equivalent
   water. For the glycoprotein and total staining methods, skip the fixing step since it was already done in
   the phosphoprotein staining method. This type of staining gives a higher background and an increase
   amount of speckling. For a more visually pleasing image of your gel, it is best to do the glycoprotein
   staining on a separate gel.

K. Imaging
   It is best to use an imager dedicated to Proteomic work. If can not do that, then you will need to protect
   your gel from keratin contamination. Therefore, you will need to place either plastic wrap or a keratin-
   free gel cutting sheet (which can be purchased through our Proteomic Center) between your gel and
   the imager. Don’t take the first piece of plastic wrap since someone may have touched it with their bare
   hands.

L. Gel and gel band storage
   Store your gel at 4°C in high quality nanopure type water for no longer than a couple days; otherwise,
   bacteria may grow. Never use parafilm to cover your gels. Parafilm leaches chemicals that interfere
   with the mass spec analysis. You will need to protect your stained gel from light. If you use aluminum
   foil, make sure there is a cover between the gel and the foil since the acetic acid in the destaining and
   storage solution will corrode the foil.

       Gel Storage Conditions:

       1) Store your gel/bands in fixing solution (10% methanol + 7% acetic acid) at 4ºC or room
          temperature.
       2) Put the band in a microfuge tube and store at -20°C until you are ready to digest.

M. Cutting gel bands
   Place your gel on a clean piece of glass, plastic wrap or a keratin-free gel cutting sheet that is covering
   the top of a uv/blue light box. Using a clean scalpel or razor, excise your band(s) of interest, making
   sure you cut as close as possible to the exact band you are interested. Don’t include neighboring,
   excess, unstained parts of the gel. If you want to be extremely cautious, you can sonicate the blade for
   5 min in a glass beaker with either acetonitrile or ethanol before cutting your band. Place the band in a
   plain clear 1.5 ml microfuge tube.

   Cut a gel slice/plug from a protein-free/unstained area of the gel so that we can check for the levels of
   keratin contamination. You will not be charged for this sample. If you are doing your own trypsin
   digestion, you will need to cut an unstained molecular weight standard band to check the efficiency of
   your digestion. Check with the facility to see what band would be appropriate.

           Band Storage Conditions (Note: you don’t need to protect your cut out band from light):

           1) Add 200 μl of 10% methanol + 7% acetic acid to each tube. Store the tube at 4°C until you
              submit it the Proteomic Center for protein identification.
           2) Store microfuge tube with the band at -20°C until you are ready to digest.

N. Submitting a band for protein identification by mass spectrometry
   Along with your bands of interest, cut a slice from an area of the gel where there should be no proteins
   as a negative control. We also need an image of your gel. Label the protein molecular weight standard
   bands and the bands that were cut out of the gel. Then fill in our sample submission form and then
   email the image and the form to masondej@ucla.edu.

O. Sending samples through the mail
   Please send the samples on ice. There is no need to send them on dry ice (save your money). Our
   shipping address is the following:

   Attn: Melissa Sondej
   UCLA Keck Proteomic Center

Dept. Chemistry and Biochemistry
       607 Charles E. Young Dr East
       MSB 1424
       Los Angeles, CA 90095-1569
       Phone: 310-206-0432

Acknowledgement
Parts of what was written above were adapted from Paul Gershon In-Gel Digestion procedure available at
http://cvr.bio.uci.edu/downloads/In%20gel%20digestion7.pdf.

Sypro Ruby Staining Protocol for Mini Gels

Day1
   1) Gently break open the gel cassette. Loosen the sides of the gel with the gel knife and cautiously
       floated the gel off of the plate into a 1000 ml pipet tip Box containing MilliQ equivalent water.
   2) Gently shake the gel and then dump off the water. Repeat the rinse a second time.
   3) Fill the Box with 100 ml Fixer/Destain Solution (10% Methanol + 7% acetic acid), and then fix the gel
       with shaking for a minimum of 1 hr.
   4) Dump off Fixer/Destain Solution into the appropriate hazardous waste bottle.
   5) Rinse the gel in MilliQ equivalent water.
   6) Dilute fresh Sypro Ruby Stain 50:50 with previously used Sypro Ruby stain to reduce the background
       and speckling. Dump 30 ml of the diluted Sypro Ruby stain into a new/another 1000 ml pipet tip Box,
       and then transfer the gel into this staining tray. If you pour the stain directly onto the gel, you will get
       background artifacts. Stain the gel overnight with shaking. (Note: protect the fluorescent dye from light
       by wrap the tray in aluminum foil. Also do not stain more than overnight since the amount of speckling
       dramatically increases.)
Day 2
   7) Transfer the gel into a clean Box containing MilliQ equivalent water. Save the used stain for later use.
       (Note: You can not use the same box you stained in unless you clean it. Otherwise the amount of
       background and speckling will dramatically increase.)
   8) Rinse the gel twice with MilliQ equivalent water as described in step 2.
   9) Destain the gel in 100 ml Fixer/Destain Solution for at least 4 hrs (towards the end of the day is even
       better because the background will further be reduced). Protect the gel from light.
   10) Immediately prior to imaging a particular gel, rinse the gel once with MilliQ equivalent water by gently
       shaking. Save the Fixer/Destain Solution for storing the gels. Replace the MilliQ equivalent water.
   11) The gel can be visualized with a laser-based scanner with an excitation ~450 nm and an emission at
       ~610 nm (FX Pro Fluorescence Imager: 532 nm excitation laser and a 555 nm LP emission filter).
       Since the Sypro Ruby stain has a second excitation maxima at ~280 nm, a blue-light or a 300 nm UV
       transilluminator can also be used without lost of sensitivity.
   12) Place the gel in Fixer/Destain Solution saved from step 10.
   13) If you would like to further reduce the background staining on the gel, continue destaining the gel
       overnight with shaking and then repeat steps 10-12. (Note: if you change the Fixer/Destain Solution
       and do longer destaining than 1 day, the faint bands may disappear.)
   14) Store the gels in Fixer/Destain Solution to prevent bacteria/mold from growing. It is best to store the
       gels at 4ºC; however, this may not always be possible. Place the gel box without foil into a Ziploic bag
       or a sealing bag. (This prevents the foil and your refrigerator from corroding.) Wrap the outside of the
       bag with foil to protect from light.

Note: SYPRO Ruby is a light sensitive fluorescence dye. Therefore, minimize light exposure after the staining
and all subsequent steps by wrapping the staining container in aluminum foil or use a black Blot Box. Careful
that the aluminum foil does not come in direct contact with the Fixer/Destain Solution or it will corrode onto

your gel. The gels can be fixed, stained and destained on either a rocking platform or orbital shaker. However,
best results are obtained with a rocking platform.

Prepared by Melissa Sondej 02/21/12