Colin Mann

La Jolla Institute for Allergy & Immunology · Division of Vaccine Discovery

I am wondering if anyone can explain some apparent weight discrepancies between SDS-PAGE and Blue Native PAGE. I understand that the underlying principle is different in each, which may produce some differences due to hydrodynamic radii. However, for example, HIV envelope trimers run on SDS-PAGE at around 100-130 kDa (ours around 100 since they're truncated). However, when the trimer is run on a Blue Native PAGE, the apparent molecular weight is around 600-700 kda. This seems universal across many publications from different groups. Taking into consideration then the hydrodynamic radius, the Env trimer is actually very similar to the Thryoglobulin ladder standard (8.1 vs 8.6). Many groups have confirmed by DLS that this 600-700 kda band is in fact trimer.

Now, based on theoretical analysis and DLS experiments, the trimer should have a total weight (including glycans) of around 350 kDa. Why then, is the trimer showing up at almost double this expected weight on NATIVE PAGE, especially when it should be well-represented by the ladder standards? Glycoproteins will migrate slower, but really this much slower?

I am most concerned with this because I am working with other viral glycoproteins, and my lab does not have DLS or another powerful method to accurately determine the weight. For RSV F trimers, for example, the SDS PAGE shows around 70 kDa for monomer, while the trimer is around 400 on Native. Given this, the expected trimer weight (~ 210 kDa) and the apparent Native PAGE weight (~400 kDa) are roughly similar in proportion to the HIV trimer. I am, however, hesitant to believe it is well formed trimers, and not, say, a dimer of trimer (aggregate).

Does anyone have any insight or advice for assessing the oligomeric form of these proteins, and possibly an explanation for this large weight discrepancy for the two types of electrophoresis?

I am having problems with some samples in SDS-PAGE. For 2 specific proteins (one viral protein one serum protein) I am getting almost consistent unexpected bands across the gel. I have included the picture. The first 6 protein lanes are 6 different constructs of the same protein on different carriers (non-reducing) the next 6 after the ladder are the same but reducing. The last two wells are NR & R of a different protein. Each was purified by a different immunoaffinity column, concentrated in a different concentrator, etc... I also run MANY other proteins day to day that do not have these bands. For some reason it is only these two unrelated proteins. I have considered uneven cooling, but that doesnt seem to make sense for this scenario. The 12 lanes should show a band around 50 and around 15 (the darkest clean bands) and the last 2 should show up at those darkest bands as well. I'm just concerned about the faint bands.

I ran it at 150V for 1 hour with slightly cooled buffer (last minute run).

Any advice/thoughts?

I am curious as to the particular mechanisms by which exogenous antibody can induce protection. Considering these antibodies do not result in natural immunity, do they have limited functions in vivo? Do these antibodies have the ability to mediate CMI by interaction with innate host cells and receptors?

For example, do passively administered antibodies solely opsonize/neutralize pathogens? Are they able to engage FcyR similarly to endogenous antibodies? Are they able to function in complement fixation or ADCC?

I have had a hard time finding this information from online sources, as most give a general definition of passive immunization without any underlying mechanisms or differences in action.

I am most interested in this in the context of immunization with immune complexes. Would co-injecting antibodies improve cellular uptake of antigens? Would this potentially stimulate development of active immunity?

Given an immunogen that does not bind germline Abs well (such as HIV GP140), could co-administering broadly neutralizing antibodies initiate adaptive responses by effecting providing T-helper cells and naive B-cells with antigen? Could this potentially enhance formation of germinal centers, which would concentrate antigen and allow for more efficient recognition by rare, antigen-binding, naive B-cells?

Thanks for the input!

I have read some papers concerning expression levels of proteins and instances of misfolding to due to "back up" during cellular processing.

I am expressing viral surface proteins and derived constructs in vitro in both 293F and Expicho. It is quite often that we will design a new construct and after purifcation, see that there is quite significant aggregation, and/or deleterious bands on the SDS-page. We often attribute this to a poor design. How likely is it that the protein is being overexpressed, causing the misfolding and aggregation? We typically add 900ug plasmid DNA per liter of transfection for 293F, and 1000ug per liter of Expicho.

During simply gravity column chromatography with either Nickel-NTA or antibody-conjugated beads, does it affect the product to have the column completely empty between steps?

I have seen some protocols say to leave a few mL of buffer on the beads between steps. Typically I allow the column to run empty (but not dry out to cracking). Does the lack of surrounding buffer affect protein integrity? I know "foamy" solutions can indicate denaturation, and was wondering if exposure to air in this way could have a similar affect, or if the lack of hydration could cause denaturation.

I know there is already several published methods, but most are low-throughput, costly and will complicate downstream GMP production.

I am trying to expedite effective cleavage of proteins by thermolysin. I have a couple of ideas, but would appreciate input on the feasibility and practicality of each.

These ideas are based on a recent work which describes a high-throughput method of Fab production by on-column cleavage of Ab by papain. I was hoping to translate to other proteolytic enzymes.

1. On-column cleavage of soluble proteins by thermolysin immobilized to sepharose

2. Traditional overnight incubation with thermolysin, but with clarified supernatant, followed by affinity chromatography to capture and elute desired protein.

3. Co-transfection with plasmid encoding thermolysin (similar to co-transfection with furin)

4. Replacement of thermolysin cleavage site with an optimized furin cleavage sequence (would this provide proper cleavage post-translationally?)

Any additional ideas would be very appreciated. I realize some of these may involve fundamental misunderstanding, which have been unsolved by research of existing literature. Any "hole-poking" would be helpful.

Thank you!

So our lab cultures about 2L of ExpiCHO a week in varying volumes from 25mL to 100mL. In the last 2 weeks we have had 4 flasks (all 100mL) show this weird growth behavior. Other transfections from the same day appear normal, and used the same media. All cultures were properly given 1st and 2nd feeds.

I personally did not perform the transfection, so issues involving human error are possible, but it supposedly was performed routinely without problems.

I am unsure as to what could even have happened, and what is exactly going on in the flask. It appears to be aggregation, and the culture looks to have grown since transfection date. Indeed while the one was a more brownish color, but have the indicative "orange" color of a viable culture, as to a pale yellow of a dead culture.

So I have run around 200 Blue native Page gels within the last year and have still been unable to figure out this issue:

When running my gels I use GE healthcare HWM Native Marker kit which is supposed to have 5 distinct bands at 669 kD, 440 kD, 232 kD, 140kD and 66kD. However, nearly every run I obtain an extra very defined band between the 440 and 232 bands. All appear well resolved as clear distinct bands. I also occasionally obtain faint bands above the 669 kD mark.

I resuspend the dried mixture in 100uL sample buffer as recommended by the manufacturer and make buffers from manufacturer stock. I run gels at 100-130V (0-15mA) for 2-3 hours.

The proteins I run on the gel always appear fine (related to protein quality of course) and typically are very sharp bands at the expected MW.

Any idea what could be the source of these extra bands could be? Is it just breakdown of the ladder standards?

I have been trying to figure out the cause of some extra bands underneath the wells of my Blue Native Page gels. I am running very high MW proteins which are intended to remain in the wells due to their size. I have accomplished this before on different occasions, but am having issue getting consistent results without the extra bands.

Does anyone have any idea why these proteins would migrate out of the well, and how I might prevent this while retaining proper separation of other samples within the gel?

I use LifeTechnologies Mini Gel Tank with NativePAGE 3-12% Tris-Glycine gels ran at 130-140V for 2.5 hours. Samples are prepared as follows:

3.5uL 4x sample buffer

0.5uL G250 loading dye

10uL protein sample in 1X PBS

10uL of sample loaded to each well.

Running buffers are prepared from Manufacturer 10X stock (cathode and anode) with cathode containing 2.5% G250 (light cathode).

Below I provided two picture. The gel from 5/29/18 is exactly how it should look. The gel from 6/25/18 shows an extra band under the wells of the analogous proteins (center 2 and ones bordering ladders).