How to Use Pipettes: A Step-by-Step Technique

Proper pipetting technique comes down to a consistent sequence of movements: setting the volume, attaching a clean tip, pressing the plunger to the first stop, immersing the tip, slowly releasing the plunger to draw liquid, and then pressing through to the second stop (the “blow-out”) to dispense. That sounds simple enough, yet small deviations in angle, speed, or depth can throw off accuracy by surprising amounts. Manual pipetting variation is a “known unknown” in laboratories, where inaccuracy and imprecision exist in practice but are rarely measured, and if those variations go unaddressed, errors compound and compromise data quality.

The Forward Pipetting Technique, Step by Step

Forward pipetting is the standard mode for aqueous solutions and the technique you should learn first. It uses the air-displacement principle: a small cushion of air sits between the piston and the liquid, and the piston’s movement controls how much liquid enters the tip. Here is the sequence, broken into its individual moves:

Set the volume: Dial or key in the target volume. If you are dialing down from a higher setting, approach the desired number from above rather than below to avoid mechanical backlash in the dial mechanism.
Attach a tip: Press the pipette firmly into a fresh tip using a slight twisting motion. A loose tip leaks air and ruins accuracy. Never touch the tip with your fingers.
Press to the first stop: Hold the pipette upright and depress the plunger smoothly until you feel the first resistance point. This is the calibrated volume stop.
Immerse the tip: Dip the tip a few millimeters below the liquid surface. Going too deep coats the outside of the tip with extra liquid; too shallow draws in air.
Release slowly: Let the plunger glide back up at a controlled, steady speed. Releasing too fast creates turbulence and air bubbles inside the tip, both of which reduce the volume you actually pick up.
Pause briefly: Hold the tip in the liquid for a second or two after the plunger returns to its resting position. This lets the air cushion equilibrate and the full volume settle.
Withdraw and move: Pull the tip out of the source liquid, touch it lightly against the vessel wall to shed any drop hanging on the outside, and move to your receiving vessel.
Dispense to the second stop: Press the plunger smoothly past the first stop and through to the second stop (the blow-out). This pushes out the last bit of liquid that would otherwise cling inside the tip.
Release away from liquid: Remove the tip from the receiving vessel before you let the plunger return. Releasing inside the liquid risks sucking some of it back up.
Eject the tip: Press the tip ejector to discard the used tip.

Throughout this entire process, keep the pipette vertical or close to it. Tilting more than about 10 to 20 degrees lets liquid creep up toward the barrel, which contaminates the internal mechanism and skews the volume you deliver.

Why Pre-Rinsing the Tip Matters

Before you start pipetting your actual samples, it helps to “condition” or pre-rinse the tip by aspirating and dispensing the same liquid two or three times. The inside wall of a dry plastic tip does not behave the same way as a pre-wetted one. Pre-rinsing coats the interior surface with a thin film of your liquid, which reduces the amount that sticks to the wall on subsequent draws. A study measuring pipetting accuracy across different matrices found that pre-rinsing the tip three times improved bias, and the improvement was especially pronounced when handling methanol and whole blood, both of which cling to plastic more aggressively than plain water.¹

Pre-rinsing is particularly important when switching between liquids with very different properties, like going from an aqueous buffer to an organic solvent. It takes only a few seconds and measurably tightens your results. If you are running a long series of the same sample type, a single pre-rinse at the start of each new tip is usually sufficient.

Reverse Pipetting for Viscous or Foaming Liquids

Forward pipetting works well for water-like solutions, but it starts to break down with viscous fluids (glycerol stocks, protein solutions, detergent-laden buffers) and liquids that foam easily. These liquids leave more residue inside the tip, and the air cushion does not behave as predictably with them. Reverse pipetting handles these situations better.

The technique flips the logic of the blow-out. You start by pressing the plunger all the way through to the second stop before immersing the tip. Then you slowly release the plunger fully to aspirate. When you dispense, you press only to the first stop, not the second. The extra volume you initially drew up stays in the tip, acting as a buffer that ensures you deliver the correct amount without having to rely on a blow-out that might splatter a viscous liquid or create foam.

The key fundamentals of both forward and reverse pipetting are considered a prerequisite for good laboratory practice, especially in biomedical research where even modest volume errors cascade through downstream assays.² If you regularly work with anything thicker than water or anything that tends to bubble, reverse pipetting is worth practicing until it becomes second nature.

Contamination Control and Filter Tips

Every time you pipette, there is a chance that aerosol droplets from the liquid travel upward inside the tip and reach the pipette barrel. Once the barrel is contaminated, every subsequent sample you handle with that pipette is at risk of carryover. This matters enormously in PCR work, radioimmunoassays, and any experiment where even trace cross-contamination can produce false results.

Filter tips are the standard defense. These are disposable tips with a small plug of hydrophobic material near the top that blocks aerosol particles from reaching the barrel. Testing with a bacterial culture showed that pipetting without a filter contaminated the pipette interior with more than a thousand colony-forming units in fewer than 100 pipetting cycles. With a high-quality filter in place, the barrel stayed sterile for over 500 cycles. The same pattern held for radioactive liquids and plasmid DNA: filters extended the contamination-free window from under 100 pipettings to 250 or more.³ When the barrel did become contaminated in unfiltered runs, carryover to fresh samples was detected in about 13% of pipetting series, which is a disturbingly high rate for any quantitative work.

Filter tips are not perfect, though. The physics of aerosol behavior means no filter can block 100% of particles across the full range of sizes. When contamination control is absolutely critical, positive-displacement pipettes offer a more reliable alternative. These pipettes use a disposable capillary and piston that make direct contact with the liquid, eliminating the air cushion entirely and the aerosol problem along with it.⁴ Positive-displacement pipettes cost more per sample, so most labs reserve them for the most contamination-sensitive applications.

How Tip Quality Affects Accuracy

Not all tips are created equal, and the interaction between the liquid and the tip surface is a bigger deal than most people realize. Standard polypropylene tips work fine for water and dilute buffers, but when you pipette complex biological fluids like whole blood, serum, or protein-heavy solutions, a surprising amount of material sticks to the tip wall. That residual film does not rinse off cleanly during dispensing, which means you deliver less than you intended and then carry trace amounts of the previous sample into the next one.

Research comparing standard untreated tips with specially treated low-retention tips showed that untreated tips left significantly more residue on their inner surface. In serial dilution experiments, the standard tips produced higher-than-expected readings at dilutions that should have been near zero, because dye from earlier, more concentrated steps was still clinging to the plastic and bleeding into later steps. Treated tips showed a steeper, cleaner drop-off in signal across dilutions and leveled off to the expected baseline much sooner.⁵

For routine aqueous work, standard tips are fine. But if your results depend on accurate handling of sticky or viscous fluids, investing in low-retention tips can meaningfully reduce both carryover and volume inaccuracy.

Ergonomics and Repetitive Strain

Pipetting looks gentle, but the repetitive thumb motions involved add up quickly. A researcher running plate-based assays can easily perform hundreds or even thousands of pipetting cycles in a single day. Biomechanical analysis of the thumb during pipetting found that all nine muscles in the thumb are actively working during each cycle. The muscle generating the highest force was the abductor pollicis brevis, the fleshy muscle at the base of your thumb. The ratio of peak muscle force to the force you actually apply on the plunger was roughly 2.3, meaning your thumb muscles work more than twice as hard as the force you feel at your fingertip. That ratio is comparable to what researchers have measured in grasping tasks, which are well-established risk factors for repetitive strain injuries.⁶

A few habits help reduce the strain:

Use electronic pipettes: Motorized plunger action eliminates thumb fatigue almost entirely. If you pipette heavily every day, this is the single biggest ergonomic upgrade.
Alternate hands: Switch between left and right hands periodically to distribute the load. Most pipettes are designed for either hand.
Take breaks: Step away from the bench for a few minutes every 30 to 45 minutes of continuous pipetting. Stretch your fingers, wrists, and forearms.
Minimize tip ejection force: Tip ejection often requires more force than the pipetting stroke itself. Some pipettes have soft-eject mechanisms that reduce this effort.
Check your posture: Keep your elbows close to your body and your wrists neutral. Reaching across a wide bench or hunching over low work surfaces amplifies strain.

Pipetting-related musculoskeletal complaints are common enough in research labs that some institutions now include pipetting technique in their ergonomic training programs, alongside keyboard and mouse use.

Using Multichannel Pipettes

When you need to fill rows or columns of a 96-well plate, a multichannel pipette handles 8 or 12 channels simultaneously, cutting the work to a fraction of what it would take with a single-channel device. The basic technique is the same as forward pipetting, but the execution is trickier because you need all channels to seal equally well and aspirate the same volume.

The most common source of error with multichannel pipettes is uneven tip seating. If one or two tips are slightly loose, those channels will aspirate less liquid or leak air. Press the pipette into the tip rack with firm, even downward force, and visually check that all tips sit at the same height before you start. When aspirating from a reagent reservoir, make sure the liquid level is deep enough that all tips submerge to the same depth. Shallow reservoirs with uneven meniscus lines will cause the outer channels to draw differently from the inner ones.

Verifying the accuracy and precision of each individual channel is important because a single underperforming channel can silently ruin an entire plate’s worth of data. Published protocols exist for assessing both accuracy and precision using manual single and multichannel pipettes.⁷ Photometric verification systems can check each channel independently using dye-based measurements, which is especially useful for catching a single drifting channel that might not show up in a bulk gravimetric test.⁸

Calibration and When to Check It

Even a perfect technique will produce wrong results if the pipette itself is out of calibration. Pipettes drift over time due to wear on the piston seal, spring fatigue, and contamination buildup inside the barrel. Most manufacturers recommend calibration checks at least once a year, though high-throughput labs or labs under regulatory oversight often check more frequently.

The two standard methods for calibrating micropipettes are gravimetric (weighing the dispensed water on an analytical balance) and photometric (measuring the absorbance of a dispensed dye solution).⁹ Gravimetric calibration is straightforward and can be done in-house with a good balance, temperature probe, and barometric pressure reading, since the density of water varies with temperature and atmospheric conditions. Photometric methods are faster for multichannel devices and can be more practical when checking many pipettes at once.

Between formal calibrations, you can do a quick sanity check by weighing 10 dispenses of water at your pipette’s nominal volume and looking at the spread. If the average weight departs from the expected value by more than a few percent, or if the individual weights vary widely, send the pipette for service.

Common Mistakes That Quietly Wreck Results

Some pipetting errors are obvious: you see a bubble, you notice you forgot to change tips, you accidentally overshoot the volume dial. The dangerous ones are the ones that look fine at the bench but show up as noise or bias in your data downstream. A few of the most frequent:

Plunger speed inconsistency: Aspirating or dispensing too fast changes the volume you actually transfer. A brisk thumb snap on the plunger might feel efficient, but it creates turbulence and can leave liquid behind in the tip. Slow, smooth strokes are more accurate.
Immersion depth variation: Going deeper into the source liquid than necessary coats the outside of the tip with extra liquid, which then gets delivered as bonus volume. Dipping just a few millimeters below the surface is enough.
Skipping the blow-out: In forward mode, stopping at the first stop instead of pressing through to the second leaves a small but consistent amount of liquid trapped in the tip. Over many transfers, this systematic under-delivery adds up.
Temperature mismatch: If the pipette has been sitting in a warm room and the reagent just came out of a cold fridge, the air cushion inside the barrel and the liquid have different thermal properties. This changes the effective volume. Letting both the pipette and the liquid equilibrate to the same temperature before starting improves accuracy.
Using the wrong pipette range: Pipettes are most accurate near the middle of their volume range and least accurate at the extremes. Using a 1000 µL pipette to deliver 100 µL introduces much more relative error than using a 200 µL pipette for the same volume. Always choose the smallest pipette whose range covers your target.

The cumulative effect of these small errors is real and measurable. In sample preparation workflows, pipetting variation can account for more analytical variability than the instrument measurement itself, which means your liquid handling, not your mass spectrometer or chromatograph, might be the weakest link in your data.

When to Consider Automated Liquid Handlers

For high-throughput work involving many samples, full well plates, and especially volumes in the low microliter range, manual pipetting becomes increasingly error-prone and physically exhausting. Automated liquid handlers can take over repetitive transfer tasks with better consistency than a human hand, particularly for long runs where fatigue and attention drift set in. Reviews of the technology note that sample preparation can account for greater analytical variability than instrument analysis, and that repetitive tasks like liquid handling benefit strongly from sensor-controlled automated systems.⁷

Automation is not a magic fix, though. Automated systems need their own calibration and verification, they require programming and validation for each protocol, and they can be expensive to acquire and maintain. For low-to-moderate sample numbers or for tasks that require frequent protocol changes, a skilled person with a well-maintained pipette is still the faster and more flexible option. The decision to automate usually comes down to volume of work: once you are routinely processing full plates several times a day, the investment starts to pay for itself in both data quality and reduced repetitive strain.

Handling Organic Solvents and Other Difficult Liquids

Water is the gold standard for pipetting because pipettes are calibrated with it, and most technique guides assume you are working with something water-like. Organic solvents like methanol, ethanol, acetonitrile, and DMSO behave differently in important ways. They are more volatile, so they evaporate from the tip during the pause between aspiration and dispensing. They have lower surface tension, so they creep more easily along the outside of the tip. And they have different viscosities, so the aspiration and dispensing dynamics change.

For volatile solvents, speed matters. Aspirate and dispense quickly to minimize evaporation losses. Pre-rinsing the tip is especially helpful here because it saturates the air column inside the tip with solvent vapor, reducing the amount of liquid that evaporates off the surface during transfer. The improvement in bias that pre-rinsing provides was demonstrated to be particularly strong with methanol.¹ For very volatile solvents or for volumes below about 10 µL, a positive-displacement pipette is often the better choice because the disposable piston makes direct contact with the liquid and eliminates the air cushion that magnifies evaporation effects.

Whole blood presents its own challenges. It is viscous, it clots, and its cellular components settle over time. Reverse pipetting works better than forward for blood. Use wide-bore tips if available, aspirate slowly, and mix the sample gently before each draw to keep the cells evenly suspended. If you are pipetting blood into analytical instruments, the carryover problem is real: proteins and cells stick to tip walls and carry over into subsequent samples unless you use low-retention tips or change tips between every transfer.

Teaching Yourself Good Technique

If you are learning to pipette for the first time, or retraining yourself after years of bad habits, the single most useful exercise is a gravimetric accuracy check. Set your pipette to a known volume, pipette deionized water onto an analytical balance, and record the weight. Repeat 10 times. Convert the weights to volumes using the density of water at room temperature (roughly 0.997 g/mL). Compare your average delivered volume to the set volume (that is your accuracy), and look at how tightly your 10 measurements cluster (that is your precision).

Do this exercise at three points in your pipette’s range: near the bottom, the middle, and the top. You will likely find that your technique is worse at the extremes. You may also discover that you have a consistent directional bias, like always delivering slightly less than the target. Once you can see the pattern, you can fix it by adjusting your speed, depth, angle, or pause time. It is the bench equivalent of recording yourself to fix your golf swing, unglamorous but effective.

1
Journal of Mass Spectrometry and Advances in the Clinical Lab. The pipetting Olympics: Propagating proper pipetting a priori in clinical LC-MS/MS analysis
2
PubMed Central. Revisiting the Micropipetting Techniques in Biomedical Sciences: A Fundamental Prerequisite in Good Laboratory Practice
3
Letters in Applied Microbiology. The effect of filters on aseptic pipetting lifetime of mechanical and electronic pipettors and carryover during pipetting
4
Nature Methods. Contamination-pipetting: relative efficiency of filter tips compared to Microman® positive displacement pipette
5
Scientific Reports. Contamination and carryover free handling of complex fluids using lubricant-infused pipette tips
6
PubMed Central. Analysis of the musculoskeletal loading of the thumb during pipetting – A pilot study
7
PubMed Central. Assessing variations in manual pipetting: An under-investigated requirement of good laboratory practice
8
JALA: Journal of the Association for Laboratory Automation. Multichannel Verification System (MVS): A Dual-Dye Ratiometric Photometry System for Performance Verification of Multichannel Liquid Delivery Devices
9
NCSL International measure. Validation of the Photometric Method Used for Micropipette Calibration