Start by including a nonionic detergent such as Triton X-100 at 0.01% in your assay buffer. This will probably suffice to block binding of reagents and compounds to the surface of the plate and improve the quality of the data.

Make sure the plate is well-mixed after starting the reaction by using a plate mixer. Be careful about the speed in order to avoid spillage.

Vmax and Km measurements will not be accurate unless you achieve near saturation with the substrate. Too low a range of substrate concentrations could be part of the problem.

Thanks so much Adam for your prompt and thoughtful reply.

To address the good points you've raised: I am achieving a nice plateau in all of my Michaelis-Menten curves, so I'm not too worried about too little substrate. I have had this problem employing both 0.6% Tween 20 in the buffer as well as with 6 mM CHAPS. I have also gotten very nice results with both of these conditions, leading me to believe this is not the key variable. Do you think triton x-100 would be better than Tween 20?

As far as mixing, do you recommend I continuously mix throughout the assay (i.e. add a 5-10 second shake step in between reads), or is mixing to homogeneity at the beginning of the reaction sufficient?

I ask because we initiate our reactions by manually adding 10 ul of 10X MgCl2 (essential cofactor) and mixing vigorously (but not so much so as to create bubbles).

Tomorrow I plan to try adding PEG-8000 (conc?) and BSA (1 mg/ml) to my buffers. I could also try a test assay with triton if you think that's better than Tween 20.

I don't think Triton X-100 is any better than Tween-20. I would say that 0.6% Tween-20 is probably too high a concentration because it is above the critical micellar concentration (CMC). If you have hydrophobic inhibitors, they will be trapped by the micelles, lowering their effective concentrations. Keep the detergent concentration a bit below the CMC.

Another consideration with detergents like Tween and Triton is their quality. Both are subject to oxidation as they age when stored in air, leading to peroxides that can damage your reagents. I always use Thermo-Fisher Surfact-Amps detergents, which come in sealed ampules under inert gas as 10% solutions. After opening an ampule, I store the unused detergent at -80. A cheaper alternative is to get a new bottle of the detergent, prepare a 10% stock solution and freeze it in aliquots. This does not apply to CHAPS, which comes as a solid.

My usual procedure is to mix the reaction only at the start. Mixing should be no more vigorous than necessary to assure homogeneity.

It seems to me that a possible source of variation in your experiment would be the quality of the nucleotide substrate, since it could be subject to degradation by DNase and RNase contamination, other than RNaseH.

Beware of interference by the inhibitor with your measurements. Check for this by running control reactions with the inhibitor but without the enzyme.

Adam, thank you so much for this information. I have explicitly tested most of the variables you mention with mostly unremarkable results, BUT I have never worried about Tween oxidation. I will pick up another bottle and store it as suggested. I have a few follow up questions:

1. If I prepare a buffer containing Tween at working concentration, say somewhere between 0.01 and 2%, I am assuming it is still vulnerable to oxidation at decreased concentration and in a pH controlled environment. Is this correct?

2. If I prepare a buffer as stated, how long would you use it if stored at room temperature, 4 C, or -20 or -80 C.

3. Will degassing my working buffer and 10% stock solution, and/or add a small

amount of TCEP, noticeably improve their shelf-life?

Feel free to skip the rest if you are willing to take my word for it that I have addressed your other concerns, but I am going to address them here in case anyone reading finds my approaches useful.

1. Tween 20 concentration is too high, and micelles are possibly sequestering our inhibitors and/or enzyme (btw, our enzyme is basically a grease ball).

a. The reason I was using such a high tween concentration is because for a long time we could only purify active enzyme in the presence of at least 1% Tween. Eventually we cracked that problem and can now produce enzyme in CHAPS below its CMC.

b. That said, I have titrated Tween 20 and CHAPS above and below their CMCs and it made no difference in any of the compounds’ IC50s that I tested.

2. Possible contamination with other nucleases either after purification or as carryover from E. coli expression and purification.

a. Yes, this is always a concern. We have had issues with contamination in the past, but they have largely been addressed.

b. We have a few pharmacological and molecular biology tools to estimate the proportion of observed activity coming from our recombinant enzyme.

i. The most rigorous but time-consuming test of contaminating activity is to purify an active site mutant which is completely dead (if pure, of course).

ii. We have one compound whose IC50s against our enzyme and the E. coli RNase H are separated by 3 logs. We use this as quick test of activity due to E. coli RNase H vs our enzyme.

iii. We also titrated cobalt against our enzyme (one that passed QC previously) and the E. coli RNase H. We found that their activity dose response curves (bell shaped) are separated by about 3 logs with no overlap on the tails of the curves. Thus, this is another test this enzyme must pass.

iv. Of course, we always use RNaseOut in our reactions which inhibits a few different RNases.

3. The compounds may be exerting non-enzyme-mediated effects on our assay.

a. I have performed the no enzyme test of compound and inhibitor with unremarkable results.

b. Additionally, I have performed IC50 assays (with enzyme) with control compounds which are identical to active compounds except for 1 atom removed to disrupt the binding motif. I have used this as an additional check for non-specific effects of a compound of similar structure in an actual reaction. Nothing of note was observed.

I'll report back when I try fresh tween and with what ultimately solves the problem for other's reference.

Again, thanks for the feedback!

Best,

Nathan

1. Yes. I would not recommend storing the detergent-containing solution for a long time, unless you freeze it. Just add the detergent from a concentrated stock solution to the buffer when you need it.

2. My usual practice with buffers is to prepare the amount I need fresh each day from a set of concentrated stock solutions (kept at 4oC), and keep them at room temperature during the day. This gives me flexibility if I want to change the composition of the buffer. For simple inorganic buffers like sodium phosphate, I just keep them at room temperature. If the buffer is difficult to prepare for some reason and can't be stored at room temp., I will store it at -20 indefinitely in 40-ml aliquots in 50 ml tubes. If the buffer is filter sterilized, you can store it (without the detergent or reducing agent) at 4oC for a long time. I don't store buffers at -80 because it takes too long to thaw them. I store the 10% detergent stock solution at -80 in 1.5-ml microcentrifuge tubes.

3. If you are asking about oxidation of the detergent, degassing the buffer and storing it under inert gas should prevent it. This seems like a lot of bother. I wouldn't take the trouble unless the buffer is unusually valuable. I don't know whether TCEP helps preserve the detergent.

Thanks for your detailed answers to my other points.