Jeffrey Perkel has been a scientific writer and editor since 2000. He holds a PhD in Cell and Molecular Biology from the University of Pennsylvania, and did postdoctoral work at the University of Pennsylvania and at Harvard Medical School.
There’s no shortage of fluorescent antibodies on the market. Between labeled primaries and secondaries, there are literally thousands available.
Researchers use these antibodies for such applications as flow cytometry, immunofluorescence, fluorescent Western blotting and more. And for many of those applications, there are catalog reagents galore from which to choose.
Sometimes, though, what’s in the catalog just won’t do. Maybe you’re working on a newly discovered protein or post-translational modification. Or studying an organism that’s not one of the traditional research models. Or maybe, you just happen to have a really good homemade antibody preparation that shows just what you want to see.
In cases such as these, if you want to tag your antibody with a fluorophore, you’re going to have to do it yourself. You can buy the reagents, of course, but kits are available, too. Here, we break down some of the key variables to bear in mind as you get going.
Choose your dye
A rose by any other name may smell as sweet, but when it comes to fluorophores, one red definitely isn’t the same as another. For all their spectral similarities, fluorophores can differ widely in brightness, photostability, hydrophobicity and more.
Such differences can have a big impact on the applications they support. Dyes that photobleach quickly are probably fine for flow-cytometry applications, as the dye need only work as long as it takes the cell to pass the laser. But if you plan to do something like confocal imaging, you’ll need a dye that’s stable under extended laser illumination.
Phycoerythrin (PE), for instance, is a relatively bright red dye that’s ideal for flow cytometric applications, says Randall Wetzel, director of cytometry at Cell Signaling Technology (CST). “If, however, a PE-conjugated antibody were used on a fluorescent microscope, it would quickly fade over a period of seconds.” Similarly, says Robin Clark, a research scientist at EMD Millipore , a company that has some 500 fluorophore-conjugated antibodies in its portfolio, APC and AlexaFluor 647 are spectrally similar; however, the former is brighter, and the latter is more stable over a range of pH values. “For intracellular applications, or in any assay where cellular chemistry is variable, one might want to take that into account.”
And then, of course, there are some more pragmatic concerns: You need a dye that is compatible with your equipment and filter sets and doesn’t spectrally conflict with other dyes you plan to use at the same time.
Choose your chemistry
Fluorophores can be conjugated to antibodies using either of two basic chemistries. NHS esters couple to amine groups (the N-terminus of the protein, as well as asparagine and glutamine residues). Maleimide chemistry targets thiol groups on cysteine residues. Of the two, NHS chemistry tends to be more popular, says Wayne Speckmann, a principal biologist at EMD Millipore, though either will work.
“With NHS [chemistry] you typically get a higher dye-to-antibody ratio than with maleimide,” says Wetzel.
Choose your ratio
On the molecular scale, antibodies are enormous relative to dyes, and it’s possible to conjugate many fluors to a single protein. But that’s not always ideal. For one thing, dye molecules can interact with one another, causing signal quenching. Furthermore, the more dyes you add per antibody molecule, the more likely you are to interfere with the molecule’s ability to enter cells and bind its target efficiently.
“If you put two dyes on an antibody, it’s going to be brighter than one. And three is brighter than two. But at some point, it actually gets dimmer as you add more dyes,” says Wetzel.
Each dye is different, says Judith Langenick, secondary antibody product manager at Abcam, and each has its own ideal dye-to-antibody ratio. For AlexaFluor 488, she recommends a ratio of four-to-eight for whole IgG, but “others are lower. And it depends on whether you want to use the product in flow [cytometry] or imaging.”
Antibody-labeling kit protocols typically provide guidance on how to achieve specific ratios. As a rule of thumb, says Wetzel, a ratio of two-to-six should work for antibodies—but the product might not be ideal. It’s entirely possible that you’ll achieve better results under different conjugation conditions.
At CST, which offers a fluorophore-conjugation service, “We perform small-scale conjugations using different dye-to-antibody ratios—for example, 4, 5, 6, 7, 8—and we test each prep to determine which is the best ratio for that antibody,” Wetzel says. Users who don’t do that, he adds, who “just do a one-shot conjugation,” may get something functional, but not necessarily ideal.
Choose your conditions
Neal Kitchen, immunoassay product manager at Thermo Fisher Scientific, used to work in technical support, fielding customer inquiries about fluorescent conjugation. One of the most common mistakes he sees is forgetting about the buffers in which antibodies are stored.
That’s because the buffers researchers tend to use for antibodies are not necessarily ideal for conjugation. Protein buffers tend to contain stabilizers like BSA, for instance, which can sop up the dye. The buffers also typically contain amine-containing compounds like Tris and azide, both of which will interfere with NHS ester chemistry. And the pH may be off, as well.
“Probably the No. 1 issue we encounter with customers … is they didn’t know the importance of the buffer not containing primary amines,” Kitchen says.
As a result, he says, Thermo Fisher—which launched a protein-labeling service earlier in 2013—always dialyzes antibodies prior to conjugation “to ensure that when we do the conjugation, we don’t have quenching from the buffer.”
You might need to purify and concentrate your antibody, as well.
Do your QC
By all accounts, one of the key tricks when conjugating fluors to antibodies is not the chemistry but the validation. “You have to be cognizant of the fact that any time you label a protein, you are potentially interfering with the function,” says Kitchen.
Thus, once you’ve concluded the labeling reaction itself, be prepared to do some QC. Use a spectrophotometer to compute the dye-to-antibody ratio. Run tests to confirm it still binds its target. And especially, compare it to unlabeled antibody in tissue-staining, flow or Western blotting (or even better, all three) to ensure that the staining pattern wasn’t changed by the chemistry.
Consider outsourcing
The fact of the matter is, conjugating fluorochromes to proteins isn’t difficult. But it does take practice, says Alejandra Solache, director of research and development at EMD Millipore. That means a lot of trial and error, wasted reagents and time and effort. (Langenick says users should expect to lose at least 10% of their antibody during the process, depending on the type and scale of the process employed. “If you want 1 mg [at the end], start with 1.1 or 1.2 mg.”)
Consequently, you might be better off outsourcing your conjugation work to an expert. Service providers generally have already done the hard work of determining optimal dye-to-protein ratios for each dye. They know the chemistry and have a wide range of dyes already available. And they can provide large batches at consistent quality—all at a price not that much higher than the kits themselves.
Joerg Pluempe, site manager at Active Motif Chromeon, says his service costs about $570 per antibody, and aliquots can be returned a week or two after receiving the antibody from the customer. (By comparison, a single Chromeo™ antibody-labeling kit from Active Motif, with enough material to label three batches of 1 mg antibody each, costs $330; IRDye® antibody-labeling kits from LI-COR cost $325 or $375, depending on scale.)
If all you need is a quick-and-dirty batch of antibody for a one-off experiment, says Kitchen, then by all means, do the work yourself. “But if you need to do more, and especially if you want QC checks, then you want to use a high-quality service provider … to make sure that the quality and QC procedures are being used every time.”
Image: Cells stained with antibodies for beta-tubulin (blue) and giantin (red). Nuclei (green) were stained with Nuclear Green. Courtesy of Abcam.