Micropipette Explained: Parts, Forward and Reverse Technique, and Accuracy Checks

A micropipette is a hand-held piston instrument that measures and transfers liquid volumes from about 1 µL to 1000 µL, and in most laboratory work it is the last thing between a planned volume and the volume that reaches the well. Small transfer errors carry through every dilution step and show up as shifted curves and poor replicates. Knowing the parts, the two techniques and how accuracy is measured separates a reliable plate from a noisy one.

This guide covers the parts of a micropipette, how to use one, forward and reverse pipetting, the ISO 8655 limits and gravimetric checks behind pipette calibration, and why peptide work adds adsorption to plastic. “How to use” here refers only to the instrument, with a generic aqueous buffer as the example liquid.

Parts of a micropipette

• Plunger. The thumb button that drives the piston. It has a rest position and two stops you can feel: the first sets the measured stroke and the second, the blow-out, travels further to clear the tip.
• Volume adjuster and display. A thumbwheel or the plunger itself turns to set the volume. When increasing, go one-third of a turn past the setting and come back down, and read the display at eye level [14].
• Tip ejector. Pushes the used tip off without touching it.
• Shaft and tip cone. The lower barrel, ending in the cone where the tip seals. A poor seal here can ruin a measurement.
• Disposable tip. The only part that touches liquid in an air-displacement pipette, and part of the measuring system: ISO 8655-2 covers tips in its scope [3], and Gilson ties its stated limits to its recommended tips [5].

The design dates from March 1957, when Heinrich Schnitger, a physician at the University of Marburg, built a prototype to spare himself hand-pipetting many small chromatography fractions [1]. Eppendorf launched the first industrially manufactured version in 1961 [2].

How a micropipette works: air versus positive displacement

In an air-displacement pipette, a cushion of air sits between the piston and the sample. When the piston rises, the air expands and atmospheric pressure pushes liquid up into the tip. Anything that changes the air, such as temperature or vapour pressure, can change the volume delivered. These pipettes are calibrated to water under defined conditions [11], and they give high accuracy and precision on aqueous, non-viscous liquids [13].

In a positive-displacement pipette, the piston is built into a disposable capillary tip and touches the liquid directly. Gilson recommends this design for viscous, dense, volatile, foaming, hot or cold liquids [13].

A multichannel pipette runs several channels at once to load plates quickly. ISO limits are doubled for these models [4], so the time saved comes with a wider tolerance.

How to use a micropipette: forward pipetting step by step

Forward pipetting is the standard technique for aqueous samples. These steps follow manufacturer guidance [14].

1. Choose the smallest pipette that covers the volume.
2. Equalize temperature. Let pipette, tips and buffer sit in the room together first.
3. Set the volume and seat the tip. Use a slight twist; do not hammer it on.
4. Pre-wet when conditions allow. Draw up and expel the set volume about three times with each new tip [4], unless temperatures differ (see below).
5. Press smoothly to the first stop, pipette vertical, tip out of the liquid.
6. Aspirate. Immerse the tip, let the plunger return smoothly, and wait about one second before lifting out [9].
7. Dispense. Touch the tip to the inside wall; press to the first stop.
8. Blow out. Pause for a second, press to the second stop, and slide the tip up the wall as you withdraw it.
9. Eject the tip and take a fresh one for the next sample.

Reverse pipetting and when it helps

Reverse pipetting draws up more liquid than the set volume, dispenses the set volume at the first stop, and leaves the remainder in the tip [12]. It is recommended for viscous, foaming, high-vapour-pressure and low-surface-tension liquids [11] [12], including buffers with enough surfactant to foam at blow-out.

The reported benefit is mostly in repeatability. In a Thermo Fisher application study, 1 µL and 10 µL transfers of 1% bovine serum albumin were repeated five times each with the pipette adjusted for each technique, and reverse pipetting cut imprecision by over 50% [15]. That is a manufacturer study with few repeats, not an independent trial.

It is not a universal upgrade. METTLER TOLEDO warns it can reduce accuracy and that precision depends on the user [12], another note cautions that it can over-deliver [22], and a 1996 article found it gave poorer inaccuracy than forward pipetting with methanol [16].

Accuracy limits under ISO 8655

ISO 8655 is the international standard for piston-operated volumetric apparatus. Part 2 sets maximum permissible errors for single and multichannel pipettes, and the Standards Council of Canada lists its 2022 edition [3]. It separates two kinds of error:

• Systematic error (inaccuracy) is how far the mean of repeated deliveries sits from the set volume.
• Random error (imprecision) is the scatter of deliveries around their own mean.

These limits at nominal volume are as reproduced in manufacturer documents [4] [5].

The limits stay fixed in microlitres across a pipette’s range [5] [6]. A 1000 µL pipette may be off by ±8 µL at any setting: ±0.8% at 1000 µL but ±8% at 100 µL [6]. The same 100 µL transfer carries a limit of ±1.6 µL on a P200 and ±0.8 µL on a P100. These figures describe the instrument under test conditions, not errors added at the bench. Sources differ at the smallest volumes (a 2 µL pipette appears at ±4.0% [4] and at ±2.5% [5]), so check the manufacturer’s specification there.

Pipette calibration and gravimetric checks

Gravimetric checking weighs what the pipette delivers and converts the mass to volume.

• Conditions. A draft-free room held within ±0.5 °C, between 15 and 30 °C, at about 50% relative humidity or above. Pipette, tips and water stand in the room for at least 2 hours first [7] [8].
• Test liquid and balance. Distilled or ion-free water, a balance whose resolution gets finer as volume falls, and evaporation control below 50 µL [7] [8].
• Replicates. Ten measurements at each of 100%, 50% and 10% of nominal volume [4] [7].
• Z factor. Each mass is multiplied by a temperature-specific factor in µL per milligram. For distilled water at 1 atmosphere, Z is 1.0029 at 20.0 °C, 1.0033 at 22.0 °C and 1.0040 at 25.0 °C [4].

A worked example, with invented numbers for illustration. A P200 set to 200 µL gives ten deliveries with a Z-corrected mean of 198.9 µL and a standard deviation of 0.5 µL. Systematic error is (198.9 − 200) ÷ 200 = −0.55%, inside the ±0.8% limit. Random error is 0.5 ÷ 198.9 = 0.25%, inside the 0.3% limit. It passes at this volume.

Between calibrations, METTLER TOLEDO’s quick check uses four replicates at 100% and 10% of nominal volume [4]. If a pipette fails, Eppendorf advises checking tip type, leaks and temperatures first; readjustment is a last resort, with all three points retested afterwards [7].

How often to calibrate

Guidance differs. Eppendorf says at least once a year, sooner with heavy use, many users or aggressive liquids [7]. Gilson suggests three to four checks a year for frequently used pipettes [14]. METTLER TOLEDO suggests monthly quick checks for pipettes used every working day and quarterly for occasional ones [4]. A reasonable approach is to set the interval by the cost of an undetected error: a pipette that defines a calibration series deserves more checks than one used for rough buffer top-ups.

In Canada, the Standards Council of Canada publishes accreditation scopes for calibration laboratories. One example, the ISO/IEC 17025 scope of Techmétrique Inc. in Saint-Lambert, Quebec, lists single and multichannel pipettes by ISO 8655 from above 20 µL up to 10 000 µL [21].

Where the errors come from

Eppendorf publishes reference values for common influences as a percentage of nominal volume [7].

Temperature and pre-wetting interact

A technical analysis co-authored with the Fraunhofer Institute for Silicate Research names the liquid-to-pipette temperature difference as the largest error source, at roughly 0.3% per kelvin at nominal volume, while the shared temperature level matters far less [9]. Skipping the pre-rinse cut that effect to about one-third to one-sixth, and hand warmth can also shift volume [9]. A German calibration service report covers these factors [10].

So pre-wetting is not automatically good. When temperatures match, it avoids a low first delivery. When they differ and cannot be equalized, BRAND advises against pre-wetting and for changing the tip after each step [11]. Eppendorf also notes systematic deviations with forward pipetting when liquid is not at 22 °C [20]. Buffer straight from the fridge is the everyday example; our peptide storage guide covers cold-stored reagents.

Angle, depth and thumb

Tilting a 1000 µL pipette by 45° during aspiration raises the volume by about 0.28% at nominal [9]. Depth is more nuanced: one note recommends about 1 cm [22], while the Fraunhofer-BRAND analysis found no direct effect and only a small indirect one, below 0.02% per millimetre on large pipettes [9]. In practice, immerse enough to avoid drawing air and stay consistent. Thumb control follows the same logic: the same smooth pressure and rhythm in every well.

Why small transfer errors matter in peptide work

A systematic error in the transfer volume compounds along a dilution series. Take a 1-in-10 series made by moving 100 µL into 900 µL of buffer. If each transfer delivers 101 µL, every step’s ratio is 1.009 times the intended value, and after five steps the last point sits about 4.6% high (our arithmetic, for illustration). Random error partly averages out across replicates; systematic error moves the whole curve. Our peptide calculator handles the arithmetic but cannot correct what the pipette actually delivered.

Quantitative methods depend on accurate standards, which is one reason analytical method validation tests accuracy and precision rather than assuming them. An area-percent purity figure from HPLC is a ratio within one run, so it is less exposed to a small volume error than a concentration result.

Peptides stick to surfaces

A perfectly delivered volume can still contain less peptide than planned. Kristensen and colleagues studied three cationic peptides (mastoparan X, melittin and magainin 2) in 220 µL of 1 µM solution, and recovered only 10–20% after 1 hour in borosilicate glass or polypropylene. Losses were smaller in Protein LoBind tubes and at higher concentrations. Passing 250 µL of 5 µM solution through four glass or polypropylene containers left recovery close to 0% [17]. A study of eight radiolabelled endocrine peptides over 48 hours concluded that container choice prevents unpredictable loss [18].

Those results are specific to the peptides tested, and they measure containers rather than tips. Amphipathic sequences such as LL-37, a 37-residue helical peptide [19], are obvious candidates for a check. Low-retention tips are a reasonable extension, not a proven fix. Keep transfers few at low concentrations and run a recovery test with your own peptide. General handling is covered in our WHMIS lab safety guide.

What’s established and what isn’t

Sourcing for Canadian labs

For labs sourcing research peptides in Canada, a few checks are worth making before an order:

• Know what you are reading. Our guide to reading a laboratory report explains what each section shows.
• Plan for stability. Review how peptides degrade before a long experiment, and see our notes on buying for a university lab.
• Compare suppliers on documents. See our research peptides in Canada overview.

GHRP-2A six-residue peptide; comparing it across a concentration series depends on accurate transfers.View product GHRP-6The related hexapeptide; side-by-side in-vitro comparisons need matched small-volume transfers.View product LL-37A 37-residue amphipathic, helical peptide, so surface adsorption is worth testing in your own tubes.View product

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