Stock → Working Solution Planner with Pipetting Steps

Different problem shape. This planner takes one stock and gives you the pipetting volumes to land on a working concentration, either in one step or in a chain of intermediates if the math demands it. The serial-dilution tool does the opposite end of the workflow: serial 10-fold transfers across a row of tubes so you can plate, count colonies, and back-calculate CFU/mL using the 30 to 300 countable rule. If you're prepping antibody dilutions, kinase buffer, or any reagent where you control the start and end concentration, you want this page. If you're titering a bacterial stock, you want serial dilution.

Three 1:10 steps in a row, not one 1:1000 in a single shot. Each intermediate sits in a tube of its own, and the planner gives you the stock volume plus diluent volume at every step. A common pattern: 10 µL of 1000× stock into 90 µL diluent to get 100×, then 10 µL of that into 90 µL to hit 10×, then 10 µL into 90 µL to land at 1×. Mix thoroughly at each stage (a quick vortex, brief spin) before you draw the next aliquot. Otherwise stratification will shift your downstream concentrations by several percent.

It's the threshold below which the planner refuses to give you a single-step plan and forces an intermediate dilution. For most labs, 1 to 2 µL on a calibrated P2 is the practical floor. Gilson's published Pipetman Classic spec on the P2 is roughly 12% inaccuracy at 0.2 µL and around 1.2% at 2 µL, which tells you why the floor matters. Set it to whatever your worst pipette delivers reproducibly with the technique you actually use, not the catalog spec. Filter tips on aqueous reagents, 1 µL is workable. Glycerol stocks or DMSO, push it to 2 µL.

Two things, and neither involves the chemistry. The first is pipette dead volume, the slug of liquid that sits inside the tip and reservoir after the last aspiration. The second is meniscus loss along the walls of the tube during dispensing, plus whatever you lose at the cap when you close it. For a single-step dilution into a 1.5 mL tube, 5 to 10% overage covers normal handling. If you're aliquoting from a reservoir into multiple destination tubes, push it to 15%. This is bench mechanics, not stability or precipitation, so it doesn't scale with reagent class the way RNP or lipid overages do.

Convert when the assay or downstream protocol expects a different unit than the label gives you. SDS at 10% w/v is fine to keep as a percent for running buffer prep, since downstream protocols call for it that way. Same with 70% v/v ethanol for surface decon. But the moment you need stoichiometry, like binding constants or enzyme kinetics, switch to molarity using the molecular weight from the Sigma-Aldrich product page. NEB usually publishes both for their enzymes. The planner supports either unit. Pick the one your endpoint protocol actually uses so you don't have to back-convert under deadline.

For dilution math, yes. For pH, you still measure. Diluting a 10× Tris-HCl buffer to 1× doesn't shift the pKa, but the buffering capacity drops by an order of magnitude, and any small contamination on the bench will move the pH further than you'd expect. The planner gives you the volumes; check pH on the diluted working solution before you commit it to a real experiment. Cold Spring Harbor Protocols has a useful entry on Tris pH temperature sensitivity (the pKa shifts about 0.03 units per °C), which matters if you're calibrating at room temp and running on ice.

You recalibrate the input, not the output. For protein stocks, drop a quick A280 on a NanoDrop or run a BCA before you start the prep, and enter the measured concentration as your stock value rather than the label value. For nucleic acid stocks, the same applies with A260. Small-molecule stocks in DMSO can concentrate slightly from evaporation across freeze-thaw cycles. If you've thawed an aliquot more than 3 or 4 times, treat the label as an upper bound and assume the real concentration is 5 to 10% higher. The planner's math is only as good as the stock value you put in.

Yes, almost always. Below 2 µL on a P2 you're in territory where tip wetting, retention, and operator technique each contribute more than 5% relative error, and they don't cancel out. The fix is a pre-dilution: take your stock, dilute it 1:10 or 1:100 into a clean tube, then prep from that intermediate. The planner's multi-step mode does this automatically once you flag the minimum volume. Cost: one extra tube and 30 seconds. Payoff: a working concentration you can defend if anyone asks why your replicates agree.

For most primary antibody dilutions you're doing 1:100 to 1:1000 from a manufacturer stock, so 15 to 20% overage covers tip retention plus the dead volume in whatever buffer reservoir you're aliquoting from. Antibodies are usually the limiting reagent by cost, so the temptation is to skimp. Resist that, because re-prepping a primary dilution at slide 11 of 12 is worse than wasting 30 µL. If you're using a multichannel for staining a whole 96-well plate, bump the overage to 20 to 25% and pre-pool the dilution in a reservoir before you start.

Three usual suspects. Pipette miscalibration is the biggest, and it's silent until you check (the lab manager who calibrates yearly is your friend). Reverse vs. forward pipetting matters too, especially for viscous stocks like glycerol-based protein storage buffers. Last one, temperature: aqueous volumes expand about 0.02% per °C, so a stock pulled cold from the freezer and dispensed before equilibration measures low. None of these are calculator problems. If the planner's math says 10 µL and you get 9.4 µL on the bench, your pipette wants service, not the math.

Whenever the single-step pipette volume drops near or below the pipette's accuracy floor. Past about a 100-fold dilution, you're typically pipetting 1 to 2 µL of stock, and the relative error climbs fast. Two 1:10 steps spread the dilution across two independent pipetting events at comfortable volumes, where each step's relative error is small. One step at 1:100 forces a tiny aspiration with much higher relative error. JoVE and protocols.io both host bench-tutorial entries on pipetting accuracy that walk through this empirically. Rule of thumb: anything beyond 1:100 from a stock should go through at least one intermediate.

The C₁V₁ = C₂V₂ math is solvent-agnostic, so the volume calculations are valid. What changes is the pipetting accuracy and the safety practice. DMSO has viscosity around 2 cP at 25 °C, roughly twice water, so air-displacement pipettes systematically under-deliver unless you slow the aspiration and pre-wet the tip. Chloroform is worse. Positive-displacement pipettes are standard practice there. Ethanol evaporates from the tip during dispense, especially at room temperature, which biases small-volume aliquots low. The planner gives you correct math. Your technique has to match the solvent. Don't transfer DMSO with a pre-set repeat dispenser unless you've validated the volumes empirically.

Right now, the planner shows a step-by-step output you can copy into your notebook or an electronic LIMS like Benchling or eLabFTW. A direct PDF export is on the roadmap. In the interim, most users either screenshot the steps or paste the volumes into the protocol field of their LIMS. If you keep paper records, the calculation summary prints cleanly from any browser using Ctrl+P, and the volumes round to two decimal places by default. Whatever export path you use, archive the input parameters alongside the output volumes, since "why did we make 100 µL at 50 µM" is a question that comes up six months later.

The dilution math is plain C₁V₁ = C₂V₂, which is universal. The pipetting accuracy thresholds come from Gilson and Eppendorf's published tolerance specs for air-displacement pipettes. For overage guidance on multichannel reservoir work, NEB's enzyme master mix protocols and Cold Spring Harbor Protocols dilution entries are reasonable references. protocols.io hosts user-contributed dilution procedures for specific reagents (antibodies, transfection lipids, small molecules) that are worth checking before you commit a large prep. The tool doesn't invent new chemistry. It keeps the bookkeeping honest and flags volumes the pipette won't handle.