# Pipetting Technique for Small Volumes (1-10 µL) Transferring 1 to 10 µL accurately is one of the most frequently performed and most frequently botched skills in the [molecular biology](/blog/careers/molecular-biology) laboratory. A single qPCR reaction, a restriction digest, a ligation, or a sequencing library prep can hinge on delivering 1, 2, or 5 µL of enzyme, primer, or template into a tube that holds the entire experiment. Get that volume wrong by 20 percent and the reaction may fail in ways that are expensive and slow to diagnose. This article covers the physical principles behind small-volume error, a step-by-step forward pipetting technique, reverse pipetting for difficult liquids, calibration and verification methods, troubleshooting, and the mistakes that trip up students and experienced researchers alike. Hands-on time to learn the core technique is roughly 20 to 30 minutes with a P10 or P2 pipette, a beaker of water, and a small analytical balance. Total elapsed time to develop reliable muscle memory is one to two practice sessions. The method applies to any air-displacement pipette in the 1 to 10 µL range and transfers directly to molecular biology, microbiology, biochemistry, and clinical bench work. ## Why 1 to 10 µL Is the Hardest Range to Pipette The problem is physical, not psychological. Every error mechanism that affects a 1000 µL transfer also affects a 2 µL transfer, but at small volumes the relative size of each error grows because the delivered volume shrinks while the sources of error stay the same absolute size or even grow. The concern about small-volume measurement accuracy is old. In 1924, Alexander Fleming published a detailed analysis of the accuracy of measuring small fluid volumes with a capillary pipette, together with a graduated pipette design for making rapid, accurate dilution series, which shows that laboratories were wrestling with this problem almost as soon as liquid handling became quantitative [1]. Modern mechanical pipetting of very small volumes dates at least to the late 1950s, when devices capable of delivering 0.1 µL down to roughly 1 nL were described for analytical work [2], and automated approaches to small-volume dispensing were being optimized decades later [3]. ### How an Air-Displacement Pipette Works A standard laboratory pipette is an air-displacement pipette. When you press the plunger to the first stop, a piston inside the handle moves down and pushes a fixed volume of air out of the tip. When you release the plunger with the tip in liquid, the piston rises and the reduced pressure inside the tip draws liquid up to replace the air that was expelled. The liquid never touches the piston. A trapped column of air, sometimes called the air cushion or dead air space, sits between the piston and the liquid surface inside the tip. The volume delivered therefore depends on the behavior of that air cushion. Air is compressible, its pressure changes with temperature, and its volume changes with humidity. At 100 µL these effects are noise. At 2 µL they are the signal. ### Where the Error Comes From Three mechanisms dominate inaccuracy at 1 to 10 µL. First, the air cushion compresses. When you draw liquid into a tip, the pressure inside the tip must fall below atmospheric pressure to lift the liquid against gravity and surface tension. The air cushion expands and its pressure drops, which means the volume of air in the tip is no longer equal to the volume of liquid drawn. The correction factor the pipette's spring and piston geometry are calibrated to compensate for is optimized at mid-to-high volumes. At 1 µL on a P10 or P20, the compression correction consumes a much larger fraction of the set volume. Second, dead volume and tip retention dominate. The film of liquid left clinging to the inner wall of the tip after dispensing, along with the droplet that never fully leaves the orifice, is a fixed volume that scales with tip geometry, liquid viscosity, and surface properties. A residual film that costs 0.3 µL out of a 100 µL delivery costs 0.3 percent. The same film costs 30 percent of a 1 µL delivery. This is the same class of problem reported in other small-volume handling contexts, where measured recovery of a biomarker drops sharply as the sampled volume shrinks because the container surface consumes a proportionally larger share of the sample. Third, technique variables multiply at small volumes. Immersion depth, tip angle, plunger release speed, and the dwell time before withdrawal all have fixed absolute consequences that become large percentages of a small set volume. A review of medicine preparation errors involving volumes under 0.1 mL in pediatric hospital practice found that administration errors were attributed to technique, calculation, dilution, and equipment problems, and that measurement of volumes under 0.1 mL accounted for a substantial share of all medicine manipulations in that setting [4]. The laboratory equivalent is direct: at sub-10 µL volumes, your hands and your equipment are both being asked for precision that leaves no slack. ## Materials and Equipment | Item | Specification or working detail | Purpose | |--|--|--| | Air-displacement pipette | P10 or P2 (1 to 10 µL or 0.5 to 10 µL range) sized so the target volume sits in the upper half of the range | Delivering the set volume | | Compatible pipette tips | Low-retention tips recommended for protein solutions, detergents, and viscous buffers | Minimize residual film | | Reagent reservoir or tubes | Sterile, low-bind if working with [nucleic acids](/blog/guides/nucleic-acids) or enzymes | Source and destination vessels | | Analytical balance | Readable to 0.1 mg (4 decimal places), draft shield closed | Gravimetric verification of technique and calibration | | Deionized water | Room temperature, equilibrated to bench conditions for at least 2 hours | Test liquid for calibration checks, 1 µL weighs 1.0 mg | | Waste beaker | For discarded blow-out volumes during reverse pipetting | Prevents dose carryover | | PPE | Lab coat, nitrile gloves, eye protection | Standard chemical and biological safety | | Forceps or tip ejector | Avoid touching tips with bare hands | Prevents contamination and moisture transfer from skin | | Calibrated balance weights | For periodic balance verification | Confirms the reference used for calibration checks | Keep the water and test tubes at ambient temperature. Cold liquid is a hidden cause of inaccuracy and is addressed in the troubleshooting section below. ## Pipetting Small Volumes Accurately Step by Step This is the standard forward pipetting technique for a clear, aqueous sample in the 1 to 10 µL range. The reason for each critical step is given so that you understand what you are protecting. 1. **Select the smallest pipette that fits the volume.** For a 3 µL transfer, use a P2 or P10, not a P20 or P1000. Pipette accuracy is specified as a percentage of the delivered volume, and that percentage widens dramatically as the set volume falls toward 10 percent of the pipette's maximum. A 3 µL set on a P20 sits at 15 percent of range, where relative error is far worse than at 3 µL on a P10. This single choice is the largest accuracy lever you control. 2. **Set the volume going up, not down.** If you need to adjust the dial, rotate past the target from below and come back up to it, or approach the final value from a lower number. Approaching from above leaves backlash in the gearing that can shift the true delivered volume. Dial smoothly and never force the micrometer past its range, which can destroy the calibration. 3. **Fit a fresh tip and seat it firmly.** Press the pipette onto the tip with a slight twist until you feel or hear it seal. An unsealed tip leaks air under the negative pressure of aspiration, and the leak is proportional to the pressure differential, which is largest during the initial draw. At 2 µL, a small leak during aspiration is a large percentage error. If you are working with proteinaceous or detergent-containing solutions, use low-retention tips at this step. 4. **Pre-wet the tip two to three times.** Aspirate and dispense the set volume into the source vessel two or three times before taking the real aliquot. Pre-wetting humidifies the dead air space inside the tip. A dry air cushion loses water vapor into itself during the first aspiration, which shrinks the cushion and reduces the delivered volume on the first draw. Pre-wetting makes the first delivery behave like the second. For very small volumes, especially with volatile or warm liquids, this step alone can remove several percent of systematic error. 5. **Hold the pipette vertically during aspiration.** Keep the pipette within about 10 degrees of vertical. Tilting the pipette changes the hydrostatic head of liquid in the tip and changes the effective volume drawn. The tilt error is worse when you hold the pipette at 45 degrees to reach into a small tube, which is exactly the habit that develops when people work with 0.2 mL PCR tubes. 6. **Immerse the tip tip 2 to 3 mm into the liquid.** Deeper immersion increases the hydrostatic pressure on the air cushion and forces extra liquid in during aspiration. Shallower immersion risks drawing air, especially if the liquid surface moves or the vessel is nearly empty. For 1 to 10 µL, 2 to 3 mm is the standard working depth. This is easier to control in a reservoir than in a 1.5 mL tube, so transfer master mixes and buffers into a reservoir when you plan to dispense many small aliquots. 7. **Press the plunger to the first stop and pause briefly.** The first stop corresponds to the set volume. Pressing to the second stop before aspiration draws extra air into the tip and gives you a liquid volume that does not match the dial. Pause about one second after reaching the first stop so the piston has fully stopped moving before you withdraw. 8. **Aspirate slowly and smoothly.** Release the plunger with steady, controlled pressure. Rapid release creates turbulence and mist inside the tip, some of which clings to the tip wall as retained liquid, and can allow bubbles to form. Bubbles at small volumes are catastrophic because a 1 mm bubble in a narrow tip orifice can represent a meaningful fraction of the set volume. If you see a bubble, dispense back to source, pre-wet again, and repeat. 9. **Pause at full draw, then withdraw.** Hold the tip in the liquid for one second after the plunger reaches its rest position, then withdraw the tip from the liquid touching against the vessel wall if you need to knock off a clinging drop. In viscous liquids such as glycerol-containing enzyme stocks, extend this pause to two or three seconds because the liquid continues rising into the tip after the piston stops. 10. **Dispense against a wall, not into open liquid.** Touch the tip to the inside wall of the destination tube at a slight angle, just above or at the liquid meniscus if there is one. Press smoothly to the first stop, pause one second, then press to the second stop to expel the residual droplet hanging in the orifice. The wall breaks surface tension and pulls the droplet off the tip. Dispensing into the middle of a liquid volume leaves a wall-clinging film and a droplet you never recover. 11. **Keep the plunger at the second stop while you withdraw the tip.** Do not release the plunger until the tip is out of the liquid and clear of the vessel. Releasing early sucks the residual film back up and can draw destination liquid into the tip, which contaminates your next aliquot and inflates the retained dead volume. 12. **Eject the tip into a proper waste container.** For one-off small-volume transfers of enzymes or master mixes, changing tips between each dispense prevents carryover. For repeated aliquots from the same source, reuse the tip deliberately and consciously, never accidentally. Practice this sequence on water until the rhythm is automatic. The full cycle from tip seating to ejection should take about eight to twelve seconds without rushing. ## Reverse Pipetting for Viscous, Foaming, or Volatile Samples Reverse pipetting reverses the stop sequence and deliberately over-aspirates so that the excess, not the residue, absorbs the retention error. The technique is the standard answer when forward pipetting fails with glycerol-rich enzyme stocks, 50 percent glycerol storage buffers, serum, detergent solutions, or volatile solvents. The sequence is as follows. Press the plunger all the way to the second stop. Immerse the tip 2 to 3 mm and aspirate the sample slowly, letting the plunger rise all the way to its rest position. The tip now contains the set volume plus a known surplus. Dispense by pressing smoothly only to the first stop. The set volume leaves the tip and the surplus stays inside. The surplus, not your sample, absorbs the residual film and the retained droplet. Eject the tip with the surplus still inside, or dispense the surplus back into a waste vessel if it is compatible. Use reverse pipetting when the liquid is viscous, prone to foaming, or when you are dispensing repeated aliquots and want to minimize between-aliquot variability. Do not use reverse pipetting with foaming liquids if the foam enters the tip, because foam breaks the liquid column and delivers air instead of sample. For repeated reverse pipetting from the same stock, change tips if any carryover would matter, because the retained surplus carries the previous aliquot's composition. ## Pipette Calibration and Performance Verification Pipette calibration is the periodic process of checking that a pipette delivers its set volume within the manufacturer's stated tolerance, and adjusting it if it does not. Even a perfect technique cannot compensate for a pipette that has drifted. Drift comes from spring relaxation, seal wear, corrosion of the piston from salts and acids, and mechanical shock from dropping the instrument. Calibration of small-volume measurement devices has been a formal concern since at least Fleming's quantitative work on capillary pipettes in the 1920s [1], and modern laboratories treat it as routine metrology rather than an optional practice. **Gravimetric verification.** The working method is to weigh what the pipette delivers. Because 1 µL of water at room temperature weighs very close to 1 mg, a 4-decimal-place analytical balance can verify 10 µL deliveries as a mass near 10 mg. The procedure is straightforward. Equilibrate a small beaker of deionized water on the bench for at least two hours so it matches ambient temperature. Tare the balance with an empty weigh boat. Pipette 10 µL of water into the boat using the full forward technique including pre-wetting. Record the mass. Repeat at least ten times. Calculate the mean and compare it to the [expected value](/blog/data-analysis/expected-value-definition-formula-examples). The spread of the individual measurements reflects your precision, and the offset of the mean from 10 mg reflects systematic inaccuracy from the pipette, the tip, the technique, or all three. **Interpreting the result.** If the mean is close to target but the spread is wide, your technique is inconsistent, and the fix is practice with attention to immersion depth, release speed, and dwell time. If the mean is consistently offset, the pipette itself may be out of calibration, the tip may be a poor match, or a systematic technique error such as inconsistent angle is present. Systematically low delivery at small volumes is most often tip retention or a dry tip, and pre-wetting is the first fix to try. Systematically high delivery often indicates aspiration past the first stop or excessive immersion depth. **Calibration schedule.** Follow your institution's policy and the pipette manufacturer's documentation for calibration intervals, which commonly involve in-house verification on a monthly-to-quarterly basis and full recalibration by a qualified service on an annual basis or after any drop, exposure to corrosive liquids, or observed performance failure. A pipette that has been dropped or that has aspirated a corrosive reagent should be checked before further use regardless of schedule. Historical and specialized literature reinforces the point that delivering very small volumes reliably requires either well-characterized instruments or purpose-built hardware, from early capillary devices for dilution series [1] through dedicated microliter and nanoliter delivery devices [2] and automated small-volume pipetting systems designed to control the systematic errors of manual work [3]. ## Controls and Expected Results A pipetting practice session should include built-in controls so you can distinguish technique error from instrument error. **Negative control.** Weigh a blank dispense in which you go through the full motion with an empty tip pressed to the second stop, or simply tare the balance, wave the empty tip over the boat, and confirm no mass appears. Any mass here means residual water on the outside of the tip, which is a withdrawal technique error. **High-volume reference.** Deliver 10 µL ten times with a P10, where the set volume is at the top of the range. This is the instrument's best case. Then deliver 1 µL ten times with the same pipette, where the set volume is at 10 percent of range. Comparing the two tells you how much of your total error is instrument range effect versus pure technique. **Cross-instrument check.** If you have both a P2 and a P10, deliver 2 µL with each. The P2 should be visibly tighter. If it is not, the P2 itself is suspect and pipette calibration is due. **Expected results.** With good technique, a healthy P10 delivering 10 µL onto a 4-decimal balance should give replicate masses clustered tightly around 10 mg, with individual readings typically within about 1 percent of the mean and the mean within about 2 percent of the target, consistent with typical manufacturer specifications for air-displacement pipettes at the top of their range. At 1 µL the same pipette will show visibly wider relative spread, and readings of 0.8 to 1.2 mg on an uncalibrated bench setup are common even among careful operators. The lesson is quantitative: small volumes tolerate less technique slop, so every source of variance you can remove, such as temperature equilibration, pre-wetting, and vertical holding, matters proportionally more. ## Troubleshooting ![Branching troubleshooting path from a wrong delivered volume to dry tip, deep immersion, fast release or cold reagent](/article-images/illustrations/pipetting-small-volumes-accurately-troubleshoot-path.webp) | Symptom | Likely cause | Fix | |--|--|--| | Delivered volume reads low on the balance, consistently | Dry tip, first-aspiration evaporation, or tip retention | Pre-wet the tip two to three times before the real draw, use low-retention tips, slow the dispense | | Delivered volume reads high, consistently | Aspirating past the first stop, excessive immersion depth, or tilted holding | Aspirate to first stop only, immerse 2 to 3 mm, hold vertical | | Wide scatter between replicates | Inconsistent release speed or dwell time, or worn pipette seals | Slow down, pause one full second at full draw and at full dispense, schedule pipette calibration | | Bubble visible in the tip after aspiration | Aspiration too fast or tip orifice partially blocked | Pre-wet, aspirate slowly, inspect the tip orifice, replace the tip if fouled | | Liquid stays hanging on the outside of the tip | Withdrawal through the liquid surface too fast or tip immersed too deep | Touch off against the vessel wall on withdrawal, reduce immersion depth | | Droplet will not leave the tip during dispense | Dispensing into open liquid instead of against a wall | Touch the tip to the destination tube wall and press to the second stop | | Volume appears correct when pipetting water but reactions fail | Cold reagents taken straight from ice, or viscous enzyme stock under-delivered | Equilibrate reagents to room temperature when the protocol allows, use reverse pipetting for glycerol-containing stocks | | Replicates drift downward across a series | Evaporation from the source during a long aliquot series, or air cushion drying | Keep the source vessel closed between draws, re-pre-wet periodically | | Volume changes when a new tip lot is used | Tip-to-pipette fit mismatch or different retention properties | Verify tip compatibility with the pipette model, re-run the gravimetric check with the new lot | | Pipette feels stiff or jerky at the plunger | Piston or seal contamination after exposure to salts or solvents | Stop using the pipette, arrange service and pipette calibration, check delivery before resuming work | ## Variations and Related Techniques **Positive displacement pipettes.** A positive displacement pipette uses a plunger-tipped capillary that directly contacts and displaces the liquid, eliminating the air cushion entirely. This removes the compression and evaporation error sources and is the preferred tool for viscous, volatile, or high-density liquids. For routine 1 to 10 µL aqueous transfers, air-displacement pipettes are standard, but keep positive displacement in mind for concentrated glycerol stocks and organic solvents. **Capillary and glass microcapillary delivery.** Before adjustable air-displacement pipettes dominated the bench, quantitative small-volume work relied on calibrated capillaries. Fleming's 1924 study examined the accuracy of capillary pipette volume measurement in detail and introduced a graduated capillary pipette for making accurate dilution series rapidly [1]. Handled microcapillaries remain useful when a fixed, known volume must be delivered repeatedly, because a capillary is calibrated once and then behaves identically on every use. **Nanoliter and drop-on-demand dispensing.** Below 1 µL, dedicated hardware takes over from hand pipetting. Nanoliter deposition systems have been built to place droplets of defined small volumes for analytical applications such as total reflection X-ray fluorescence sample preparation [5], and drop-on-demand dispenser systems have been evaluated for dosing small volumes for chromatographic quantification [6]. Hand-pipetted volumes of 1 to 10 µL sit above this range, but knowing where the hardware boundary lies clarifies why your bench pipette was never designed to deliver 200 nL accurately. **Automated liquid handlers.** Automated pipetting systems designed for small volumes were being described and refined in the 1990s as a way to control the systematic and random errors of manual small-volume work across many samples [3]. For plate-based workflows in which dozens or hundreds of 1 to 10 µL transfers are needed, automation removes operator variance, though the same physical principles, such as pre-wetting and tip retention, still apply to the machine's tips. **Application contexts that live in the 1 to 10 µL range.** Small-volume handling shows up wherever sample is scarce. Multiplex bead-based immunoassays have been developed specifically to quantify many antibodies simultaneously in small volumes of plasma, precisely because conventional plate assays consume more sample than is available [7]. The same constraint drives qPCR, single-cell workflows, and precious clinical specimens. In each case, the transfer technique described in this article is the limiting skill. ## Storage and Stability Notes Pipettes themselves have storage requirements that protect calibration. Store air-displacement pipettes vertically on a stand with the plunger fully released, so the spring is unloaded and the piston seal is not compressed during storage. Never lay a loaded pipette on its side with liquid in the tip, because liquid can wick into the shaft and corrode or foul the piston, which is one of the classic causes of a pipette failing its next calibration. After working with salts, nucleic acids, or corrosive reagents, wipe the nose cone and check that no liquid entered the shaft. Tips should be stored in sealed boxes or sterile racks to prevent dust and moisture pickup. Wet or contaminated tips alter retention behavior, and dust on a tip orifice can break the liquid column during aspiration at 1 µL. For the reagents you are pipetting, the small-volume context creates its own stability risks. Repeated withdrawal of 1 to 2 µL aliquots from a stock tube means many openings, each of which exposes the contents to room temperature and evaporation. Keep stock tubes on ice or in a cold block while working, except when the manufacturer specifies room temperature, close them between draws, and consider single-use aliquots into low-bind tubes for enzymes and master mixes that will be accessed repeatedly. Glycerol-containing enzyme stocks stored at minus 20 degrees Celsius are stable for months, but each freeze-thaw cycle and each extended warm handling session degrades them, so the less you open the tube, the longer the reagent performs. ## Common Mistakes and Limitations The following mistakes account for most failed small-volume transfers observed on a teaching bench. Each is stated as a specific, fixable behavior. **Using the wrong pipette size.** Setting 2 µL on a P20 instead of a P2 or P10 is the single most common accuracy error. The relative error specification of a pipette deteriorates sharply below about 10 to 20 percent of its maximum volume. Match the pipette to the volume every time. **Skipping the pre-wet.** The first aspiration from a dry tip delivers measurably less than subsequent aspirations because the dead air space absorbs vapor. Researchers who pre-wet religiously at 100 µL often skip it at 2 µL, which is exactly backwards. **Aspirating to the second stop by habit.** Forward pipetting requires the first stop for aspiration. Players who learned on a P1000 and press to the second stop out of habit will over-deliver small volumes consistently. **Releasing the plunger too fast.** Fast release aerosolizes liquid inside the tip and leaves a mist coating on the wall that never dispenses. Slow, deliberate release is the habit that separates consistent operators from erratic ones. **Ignoring temperature.** Aspirating a cold liquid, such as an enzyme stock straight from ice, into a warm pipette and warm air means the air cushion warms and expands after the draw, pushing extra liquid out, so you under-deliver. Equilibrate reagents to room temperature whenever the protocol permits, and if you must pipette cold liquid, use a positive displacement pipette or accept and pre-wet against the systematic error. **Reusing tips across incompatible solutions.** Tip carryover at small volumes is proportionally huge. Residual 0.2 µL of a stock enzyme solution in a tip can contaminate an entire downstream series. Change tips deliberately whenever carryover would matter. **Checking volume by eye.** You cannot verify a 2 µL delivery visually. A droplet that looks right in a tube can be 1.4 or 2.6 µL. Use the balance for any transfer that matters, at least periodically, and use the gravimetric check to qualify your own technique, not just the instrument. **Treating pipette calibration as optional.** A pipette that passes at 100 µL can fail at 1 µL because the error sources scale differently. Verify performance at the volumes you actually use, not only at the volume the service contract tests. **The limits of technique.** Good technique fixes operator variance. It cannot fix a worn seal, a bent shaft, or a fundamental mismatch between the liquid and the mechanism. The review of small-volume preparation errors in clinical practice reached the same conclusion from the medical side, identifying equipment limitations and technique as parallel contributors and noting that some small-volume measurement problems are best solved by avoiding the measurement altogether, for example through standardized pre-diluted stocks that keep the pipetted volume in a comfortable range [4]. On the bench, the analogous move is diluting a reagent so you can pipette 10 µL instead of 1 µL, which converts a hard small-volume problem into an easy larger-volume one. **A limitation of the skill itself.** Even with excellent technique and a calibrated pipette, the 1 to 10 µL range carries irreducible random error that grows as the volume falls. For volumes where your own gravimetric checks show unacceptable spread, change the workflow, either by scaling the reaction up, pre-diluting the reagent, or moving to a positive displacement or automated solution, rather than trusting hope. **Safety note.** Nitrile gloves protect you and the sample, but they do not substitute for engineering controls. Pipette volatile organic solvents and anything with harmful vapor inside a certified chemical fume hood. Treat all biological samples as potentially infectious, decontaminate pipette handles and nose cones with an appropriate disinfectant after work with biological material, and never mouth-pipette. Sharps and broken glass capillaries, if you use them, go into a sharps container, not the bench trash. ## Frequently Asked Questions ### Why is my pipette inaccurate below 2 µL? Air-displacement pipettes lose relative accuracy at low set volumes because the compressible air cushion, tip retention, and evaporation effects stay the same absolute size while the delivered volume shrinks. Every one of these fixed errors becomes a larger percentage of a 1 µL transfer than of a 10 µL transfer, so the specified tolerance of the instrument widens near the bottom of its range. ### Should I use the P2, P10, or P20 for a 5 µL transfer? ![Path for picking P2, P10 or P20 and forward or reverse pipetting for a small volume](/article-images/illustrations/pipetting-small-volumes-accurately-pipette-choice.webp) Use the P2 or P10. The rule is to pick the smallest pipette whose range comfortably includes the volume, ideally with the set volume in the upper half of the range. A 5 µL setting on a P20 sits at 25 percent of range, where relative error is worse than at 5 µL on a P10, which sits at 50 percent of range. ### What is reverse pipetting and when should I use it? Reverse pipetting over-aspirates by starting at the second stop and dispensing only to the first stop, leaving a deliberate surplus in the tip to absorb the residual film. Use it for viscous liquids such as glycerol-containing enzyme stocks, for foaming samples, and for repeated aliquots where forward pipetting gives inconsistent results. ### How often should pipette calibration be done? Follow your institution's policy and the manufacturer's documentation, which typically means in-house gravimetric verification on a monthly-to-quarterly basis and professional recalibration annually or after any drop or corrosive exposure. Verify performance at the small volumes you actually use, because a pipette can pass at large volumes and fail at 1 µL. ### Do low-retention tips help with small volumes? Yes, particularly with viscous, proteinaceous, or detergent-containing liquids where a surface film clings to the tip wall. Because the retained film is a fixed volume, eliminating part of it has a proportionally larger benefit at 1 µL than at 100 µL. Low-retention tips are cheap insurance for the 1 to 10 µL range. ### Can I pipette a cold sample straight from ice? You can, but expect under-delivery, because the cold liquid cools the air cushion, which then warms and expands after the draw and pushes liquid back out. Equilibrate reagents to room temperature when the protocol allows, or use a positive displacement pipette when it does not. ### Why does liquid stay in the tip when I dispense? Residual liquid indicates either a surface film that needs the second-stop blow-out against a vessel wall, or a liquid property such as high viscosity or surface tension that exceeds what forward pipetting can handle. Touch the tip to the destination wall, press fully to the second stop, and if retention persists, switch to reverse pipetting or low-retention tips. ### What is the best way to check my own pipetting accuracy? Run a gravimetric check on a 4-decimal analytical balance. Weigh at least ten deliveries of 10 µL of room-temperature water, where 10 µL should read about 10 mg. The mean tells you about systematic error from the pipette or technique, and the spread tells you about your precision. Repeat at 1 µL to see how much harder the bottom of the range is. ## Sources 1. [On the Accuracy of Measurement of Small Volumes of Fluid with a Capillary Pipette: Incorporating a Description of a Graduated Pipette for Rapidly and Accurately Making a Series of Dilutions of a Fluid](https://www.semanticscholar.org/paper/7f949ccf24f6e89c375f914d656e657494bde5ad) 2. [A PIPETTING DEVICE FOR VOLUMES OF 10$sup -4$ TO 10$sup -9$ MILLITERS](https://www.semanticscholar.org/paper/9cd6e71aeadf7ad7bf6dc7b9db819f8fc452784e) 3. [Automatic pipetting small volumes.](https://www.semanticscholar.org/paper/7199dec35dfad623eb525b452eaac28f10a04c1b) 4. [Accuracy of intravenous and enteral preparations involving small volumes for paediatric use: a review](https://pubmed.ncbi.nlm.nih.gov/31156991/) 5. [Nanoliter deposition unit for pipetting droplets of small volumes for Total Reflection X-ray Fluorescence applications](https://doi.org/10.1016/J.SAB.2013.01.006) 6. [The dosage of small volumes for chromatographic quantifications using a drop-on-demand dispenser system](https://pubmed.ncbi.nlm.nih.gov/17549459/) 7. [A semi-automated multiplex high-throughput assay for measuring IgG antibodies against Plasmodium falciparum erythrocyte membrane protein 1 (PfEMP1) domains in small volumes of plasma](https://pubmed.ncbi.nlm.nih.gov/18549480/)