Viral Proteins Are Not Migrating Down the Gel - (Jun/19/2014 )
1- Magic Mark Standard
2- WtF 6hr
3- WtZ 6hr
5- F1R 6hr
6- Mock infected
7- WtF 9hr
8- WtZ 9hr
9- RGV0 9hr
10- F1R 9hr
11- Mock infected
GAPDH probe of above membrane
I am isolating viral (Sendai virus) proteins between 4 different virus variants at 4 different time points (1, 3, 6 & 9 hr p.i.) (the pictures above are of 1 and 3 hours only) of infection. I use the LLC-MK2 cell line for my infections. After visualizing my membrane, there seems to be a lot of viral protein stuck at the top of the lane (of the gel) which did not migrate down.
My PI and I suspect that it is whole (non-disrupted) viral particles which have attached to the wells of the gel preventing migration down the lane.
Does this sound plausible? And if so, what is the best way to break up the viral particles in order to get their proteins to migrate down the gel?
-For more information see my Western Blot protocol below.
Western Blot Protocol:
To measure levels of viral protein expression, cell pellets from virus infected culture were resuspended in 200μl radioimmunoprecipitation buffer (RIPA, 50mM Tris-HCL pH 7.4, 150mM NaCl, 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS) (Boston Bioproducts, Ashland, MA) including a protease inhibitor cocktail (Thermo Scientific, Waltham, MA), then incubated on ice for 30 minutes, with vortexing every 10 minutes. The protein lysate was centrifuged at 14000 Xg for 15 minutes at 4°C and the supernatant (protein lysate) was transferred to a new 1.5 ml tube. Equivalent volumes of protein lysate and 2x sample buffer (4% LDS, 0.8M Triethanolamine-Cl pH 7.6, 4% Ficoll-400, 0.025% Phenol Red, 0.025% Coomassie Brilliant Blue, 2mM EDTA (Expedeon, San Diego, CA)) were mixed. The protein samples (protein lysate and 2x sample buffer) were denatured in a 70°C water-bath for five minutes.
Expedeon’s Run Blue Protein Electrophoresis, Dual Run & Blot Unit instruction manual was used to run the gel and blot the proteins. Briefly, the protein samples were separated by electrophoresis on a 10% SDS-polyacrylamide gel (Expedeon, San Diego, CA) for one hour at 150V. A Pre-Stained protein ladder (EZ-Run Pre-Stained Rec Protein ladder, Fisher, Pittsburgh, PA) and a western protein ladder (MagicMark XP Protein Western Standard, Invitrogen, Carlsbad, CA) were run alongside the protein samples. Tris- Tricine SDS (1.2M Tris, 0.8M Tricine 2% SDS, 50mM Sodium Bisulfite pH 8.2, Expedeon, San Diego, CA) run buffer was used. Following electrophoresis, the gel was transferred onto a nitrocellulose membrane (Whatman, Piscataway, NJ) for 1.5 hours at 200V at room temperature with ice packs and a stir bar. To prevent nonspecific binding of antibody, the membrane was blocked in 5% non-fat dry milk (Bio-Rad, Hercules, CA) in TBST (Tris buffered saline (Boston Bioproducts, Ashland, MA) and 0.1% Tween). After blocking, the membrane was incubated in a 1:2000 dilution of a rabbit anti-SeV primary antibody (MBL International, Woburn, MA) solution for one hour at room temperature with rocking, followed by washing three times, five minutes each, with TBST at room temperature. To detect primary antibody attachment to SeV proteins, a 1:1500 dilution of horseradish peroxidase-conjugated anti-rabbit IgG secondary antibody solution was added to the membrane for one hour at room temperature with rocking; followed by membrane washing with TBST three times, five minutes each, at room temperature. Proteins were visualized using the Pierce™ ECL Western Blotting Substrate kit (Thermo Scientific, Rockford, IL) and analyzed using the Molecular Imager VersaDoc™ MP Imaging System with the Quantity One Analysis Software (Bio-Rad, Hercules CA).
In order to normalize the blot results, the membranes were stripped (62.5mM Tris-HCL pH 6.8, 2% SDS, 100μM β-mercaptoethanol) at 50°C for 30 minutes and then washed five times in TBST for five minutes at room temperature. The membranes were incubated overnight at 4°C in 5% non-fat dry milk to block non-specific binding sites. On the following day the membranes were incubated with a 1:1000 dilution of mouse anti-GAPDH primary antibody (Research Diagnostics Inc, Flanders, NJ) probe for one hour at room temperature with rocking. The membranes were washed three times in TBST for five minutes at room temperature with rocking and then incubated in a 1:3000 dilution of horseradish peroxidase-conjugated anti-mouse IgG secondary antibody (Amersham Life Sciences, Piscataway, NJ) for one hour at room temperature with rocking. Proteins were visualized as described above.
Thanks for reading and any help is much appreciated!
Try boiling the lysates - 70 might not be fully denaturing. You could also try lysing your infected cells directly in the PAGE loading buffer.
Thanks for the reply! I'm taking your first suggestion and heating my samples at ~95C.
Excited to see my results!
Top: 6 and 9 hours p.i.
Bottom: GAPDH probe of the 6 and 9 hours p.i. (the stripping didn't work as well)
So I heated my samples just under boiling (bubbles formed all over the beaker but did not detach from the surface) for 5 minutes and proceeded with the same protocol stated above.
It looks like the heat increase may have helped some but there is still a lot of protein that appears to be stuck in the well.
I know you mentioned to try lysing my infected cells in PAGE loading buffer; the only problem is that I have already lysed all my infected cell pellets in RIPA buffer and have made aliquots of all my samples. As I would like to avoid infecting cells and isolating the protein again (this took a VERY long time) is there anything else I can do to my current samples such as adding a few microliters of lysis buffer?
Or should I try actual boiling water with increased time?
Boiling for extended times doesn't usually add anything, other than to cause your proteins to aggregate.
You should add 150 mM DTT or 300 mM 2-mercaptoethanol to your loading buffer and then boiling - this is there to reduce the di-sulphide bonds between cysteines and should help maintain your proteins in a linear state.
You should also spin down your boiled samples (1 min at 12,000 RCF should work) to remove protein aggregates that could be blocking your wells and inhibiting migration.
70C for 20 minutes should properly denature the virus particles under normal conditions
but, the ripa contains 1% np-40. np-40 and triton will strip sds from proteins (or prevent proper binding). you should add extra sds to the loading buffer to compensate for the presence of the np-40.
So it looks like the boiling really does help!
I tested boiling times (5, 10, 15 and 20 minutes) and the effects of the addition of Lysis buffer (5ul, 10ul and 15 ul) at 70°C and boiling for 5 min with 5ul of Lysis buffer. It looks like boiling for 15 or 20 minutes works! There seems to be a pretty high band seen in most all lanes; but not seen in lane 8. Do you think we don’t see it because of the long (20min) boiling?
Any interpretations of these results will be very much appreciated!
how long was the 70C incubation? why didn't you show 70C incubation without lysis buffer added (keep in mind that adding lysis buffer changes the final volume and dilutes the loading buffer)?
the 20 minute boiling may have eliminated the highest band but you are also seeing a reduction in (what i think is) the band of interest. based on this result, i wouldn't boil for more than 10-15 minutes (which still concerns me regarding possible aggregation).
I incubated my 70C samples for 5min. I know I should have included at least one 70C without lysis buffer, I wanted to conserve my samples; but I basically already know what happens with just a 70C incubation (my initial post).
For my purposes I need to be able to see all virus protein bands but because the time points are relatively early (1, 3, 6, & 9hr p.i.) we are unsure how many proteins may be present. For my project (Masters Thesis), I am trying to figure out a molecular mechanism responsible for the reduced replication rate of one of the virus variants (F1-R).
I'm thinking the 15 min boil will do the trick!
Thanks for all the help!!