# Pipette Cleaning Solutions Compared The best cleaning solution for pipettes is the one matched to the contaminant you are actually removing: mild detergent and distilled water for routine residue, 70% alcohol for fast disinfection between tasks, and dilute bleach when biological decontamination is the goal. No single liquid does every job well, because a pipette confronts you with three different cleaning problems at once, which are removable residue on the outside, biological contamination on contact surfaces, and salts or sample fluid that creep into the shaft and onto the piston. The rule of thumb is simple. Start with water, escalate to detergent only when water fails, reserve alcohol for disinfection rather than deep cleaning, and use bleach or specialized decontamination solutions only when the hazard demands it. This article walks through each of the best cleaning solution options for pipettes with their pros and cons, then compares them on the criteria that decide real bench choices. ## Why the Cleaning Solution You Choose Matters A pipette is a precision instrument, not a piece of glassware. The plunger mechanism, piston, seal, and O-rings inside the shaft are machined to tolerances measured in micrometers. When the wrong solution enters the barrel, it can dissolve lubricant, swell seals, corrode the piston, or leave its own residue behind. Any of those outcomes shifts delivered volume, and shifted volume only shows up later as failed pipette calibration or as unexplained spread in your assay data. Cleaning solution choice also matters for a second reason that people notice less often, which is carryover. A contaminated pipette or tip transfers traces of the previous sample into the next one. In routine cloning this may be invisible. In trace analytical work it is fatal to the result. Studies of ultra-trace contaminants in complex matrices, such as bisphenol analysis in canned condensed milk and breast milk by HPLC-DAD and LC-QQQ-MS, show how demanding sample preparation becomes when analytes sit at nanogram per milliliter levels and the matrix itself is messy [1]. Pipetting equipment that carries residue from a previous extraction is exactly the kind of hidden variable that destroys work at that sensitivity. Finally, cleaning matters because it is the step that keeps calibration honest. A pipette that is cleaned on a schedule drifts less between calibration events than one that is cleaned only when something goes visibly wrong. ## Three Different Jobs Are All Called "Cleaning" ![Decision path that starts from the contaminant type and leads to water, detergent, alcohol, bleach or a service kit.](/article-images/illustrations/best-cleaning-solution-for-pipettes-solution-choice.webp) Before comparing solutions, separate the jobs, because the best cleaning solution for pipettes changes with each one. **External decontamination.** Wiping the outside of the body, the nose cone, and the tip ejector. This removes drips, aerosol deposits, and fingerprints. Most cleaning happens here, and most damage happens here too, when aggressive liquids run into the shaft. **Internal maintenance.** Flushing or wiping the shaft, piston, and seal area after liquid has entered the barrel, which should never happen but does. This is the job where solution choice has the highest stakes, because the piston and seals are the parts that define accuracy. **Sample-contact surfaces.** Glass capillaries, patch-clamp pipettes, Pasteur pipettes, and reusable glass syringes. These tolerate far more aggressive treatment than the internal parts of a mechanical pipette because they have no piston or seal to damage. One vocabulary note prevents a common confusion. In analytical chemistry, the word "clean-up" usually refers to cleaning up a sample extract, not cleaning the instrument. QuEChERS clean-up, for example, uses sorbents such as Z-SEP, Z-SEP Plus, PSA, C18, chitin, and EMR-Lipid to strip matrix components from an extract before chromatography [1]. Similarly, a classic clean-up step for veterinary drug residues passes an extract through a small aluminum oxide column held in a Pasteur pipette before HPLC analysis [2]. Those procedures are about purifying samples. Nothing in them replaces cleaning the pipette itself, and the two activities use completely different reagents. ## Pipette Cleaning Solutions Compared: The Core Table The table below is the heart of this comparison. Contact times and concentrations shown are standard laboratory practice, and manufacturer documentation for your specific pipette always takes precedence. | Solution | Best for | Typical contact time | Material compatibility | Key limitation | |--|--|--|--|--| | Distilled or deionized water | Rinsing after every use, removing water-soluble salts and buffers | Immediate rinse, then wipe dry | Compatible with everything | Does not remove organics, grease, or microbes | | Mild laboratory detergent (dilute, roughly 0.1 to 1% working solution) | Routine external cleaning, removing dried residue and oils | Several minutes of contact, then thorough rinse | Safe on most housings and glass, keep out of the barrel | Must be rinsed completely or it leaves its own residue | | 70% ethanol | Fast disinfection of exterior surfaces between samples and at day's end | 30 seconds to a few minutes of wet contact | Safe on most housings, never inside the shaft | Poor at removing protein or salt deposits, flammable | | 70% isopropanol | Same disinfection role as ethanol, slightly better grease tolerance on exteriors | 30 seconds to a few minutes of wet contact | Safe on most housings, never inside the shaft | Can dry and stiffen rubber seals with repeated heavy use | | 10% household bleach (diluted to about 0.5 to 0.6% sodium hypochlorite) | Biological decontamination after spills, RNA work, and infectious materials | Full wet contact, typically at least 10 minutes for disinfection, then water rinse | Hard plastics and glass tolerate it, metal pistons corrode on repeated exposure | Corrosive to metal, must be rinsed off, inactivated by organic matter | | Commercial RNase and DNase decontamination sprays | Work surfaces and pipette exteriors before RNA or low-copy DNA work | Per manufacturer instructions, often a minute of contact then wipe | Formulated for lab plastics, verify against your pipette model | Cost per use, and some leave residue that must be rinsed | | Ultrasonic cleaning with detergent, applied to glass capillary tips | High-throughput reuse of glass patch-clamp pipettes | Minutes per cycle in a sonication bath | Glass and metal only, not mechanical pipette bodies | Requires equipment and validation for the application | | Manufacturer-approved pipette cleaning solutions and lubricants | Internal service during seal and O-ring replacement | Per manufacturer protocol | Guaranteed by the manufacturer for that pipette | Model-specific, and usually part of a full service rather than routine cleaning | ## Option-by-Option Review: Pros and Cons ### Distilled Water Water is the most underrated cleaning solution in the laboratory. Salts, buffers, most media, and most aqueous reagents dissolve in it instantly, and it leaves nothing behind if you use distilled or deionized grades. Tap water is a poor substitute because dissolved minerals redeposit as scale inside the nose cone and on the piston. The pros are total material compatibility, zero cost, zero toxicity, and zero residue. The cons are equally clear. Water does nothing to proteins, oils, nucleases, or microbes, and dried deposits may need detergent or a soak rather than a quick rinse. Water is the default first rinse and the mandatory final rinse after almost every other solution on this list. ### Mild Laboratory Detergent A dilute solution of a laboratory-grade detergent is the workhorse for routine cleaning. It lifts dried buffer crusts, cell culture medium, oils from handling, and general grime from the housing, hanger, and ejector. Keep the solution dilute, work with a lint-free cloth or soft brush, and rinse with distilled water until no suds remain. The main pro is that detergent actually removes things water cannot. The main con is residue. A detergent film on the nose cone transfers into your next tip and into your next sample, so the rinse step is not optional. Avoid household dish soaps with lotions, fragrances, or abrasives, because moisturizers leave films and abrasives scratch the housing. ### 70% Ethanol and 70% Isopropanol Alcohols are the standard disinfectants for bench surfaces and pipette exteriors. The counterintuitive reason 70% works better than higher concentrations is that water slows evaporation and lets the alcohol penetrate cells before it denatures their proteins. Straight ethanol evaporates too fast to disinfect reliably and can even fix organic material onto the surface. Use alcohol by spraying or wiping the outside of the pipette, the nose cone, and the ejector, and let the surface stay visibly wet for a meaningful contact time rather than wiping it dry instantly. The pros are speed, availability, evaporation without residue, and broad antimicrobial activity. The cons are that alcohol does not clean in the detergent sense, that it is a poor choice against dried protein or salt, that it must never be flooded into the shaft where it strips piston lubricant, and that repeated heavy wiping with isopropanol can dry out elastomer seals over time. A practical compromise many labs adopt is alcohol for the between-tasks wipe-down and detergent plus water for the scheduled deep clean. The two do different jobs and they complement each other. ### Dilute Bleach Sodium hypochlorite is the strongest routinely available decontaminant on this list and the standard answer for biological spills and post-RNA work. Household bleach diluted 1 to 10 in water gives roughly a 0.5 to 0.6% sodium hypochlorite solution, which is the working concentration most biosafety manuals describe. The pros are inexpensive, broad, and reliable killing of bacteria, viruses, and mycoplasma, plus chemical destruction of [nucleic acids](/blog/guides/nucleic-acids), which is why it is favored around RNA work. The cons demand respect. Hypochlorite corrodes stainless steel pistons and springs on repeated contact, it is inactivated by organic load so visibly dirty surfaces need cleaning before disinfection, it degrades plastics and coatings with long exposure, and it must be rinsed off with distilled water afterward. Wipe the exterior with 10% bleach when the situation calls for it, never pour it through the shaft, and follow with water and then a dry wipe. ### RNase and DNase Decontamination Sprays Commercial nuclease-decontamination formulations exist because RNases are extraordinarily stable and survive alcohol and many disinfectants. These sprays are designed for the exteriors of pipettes, racks, and work surfaces before RNA extraction, single-cell work, or low-copy-number PCR setup. Follow the product's stated contact time, wipe, and rinse with water when the instructions call for it, because some formulations leave a residue that can interfere with downstream enzymatic reactions if it dries on a surface that contacts samples. The pros are documented nuclease destruction and convenience. The cons are cost, the residue question, and the fact that a spray does not replace the mechanical removal of dried residue. ### Ultrasonic Cleaning With Detergent Ultrasonic cleaning occupies a specialized but important niche. In electrophysiology, patch-clamp glass pipettes have traditionally been discarded after a single recording attempt, which limits throughput badly because many attempts fail. An automated cleaning method developed for this problem sonicates the pipette tip directly within the bath solution where the cells sit, using detergent, and avoids contaminating the bath or the internal solution of the recording pipette with detergent. The published evaluation showed that patch pipettes could be reused consecutively at least ten times, that the sonication cleaning process did not harm the brain slices or other neurons being patched, and that combining this approach with automated patch-clamp substantially improves recording throughput [3]. The lesson generalizes with limits. Ultrasonic energy plus detergent is extremely effective at stripping residue from glass and hard metal surfaces, and it scales across many items at once. The limitations are equally real. A mechanical air-displacement pipette is not a glass capillary, and ultrasonicating a fully assembled pipette forces liquid past the seals and destroys the lubrication and calibration. If you use sonication for sample-contact glassware, disassemble what can be disassembled, protect what cannot, and validate the cycle for your own application. ### Manufacturer-Approved Solutions and Service Kits Every major pipette manufacturer publishes a maintenance protocol for its own instruments, naming which solutions are safe on the housing, which grease to use on the piston after cleaning, and which parts tolerate autoclaving. These protocols belong in the comparison because they are the only solutions with a compatibility guarantee for the inside of your specific instrument. The con is obvious, which is that they are model-specific and usually tied to a full service with seal replacement rather than a quick bench wipe. Keep a copy of the maintenance section of your pipette's manual at the bench, because it answers compatibility questions faster than any general rule. ### What Not to Use Acetone and other strong solvents attack many pipette housings and dissolve the lubricant inside the shaft. Concentrated acids and oxidizers pit metal and embrittle plastics. Abrasive powders scratch housings and create crevices that harbor contamination. Chromic acid cleaning solutions are a legacy approach for stubborn organic residue on glassware that no modern lab should apply near a mechanical pipette, given their extreme hazard and the availability of safer alternatives. Strong alkali such as concentrated sodium hydroxide damages aluminum components found in some pipette bodies. When detergent and water fail, the correct escalation is disassembly and mechanical cleaning with manufacturer-approved products, not stronger chemistry. ## How the Options Compare on the Criteria That Decide Real Choices **Match to contaminant.** This criterion dominates all others. Aqueous salts yield to water. Dried medium and oils yield to detergent. Microbes yield to alcohol, bleach, or nuclease sprays depending on the organism and the nucleic acid requirement. Protein crusts respond to detergent soak far better than to alcohol. Chemical residues respond to the solvent that dissolves them, which requires knowing your reagent. **Material compatibility.** Glass and stainless steel tolerate almost everything on the list. The vulnerable parts are the elastomer seals, O-rings, lubricant, and painted or coated housing. Alcohol and bleach are the two common solutions most likely to cause cumulative damage, alcohol by stripping internal lubricant when it gets inside, bleach by corroding metal on repeated exterior contact. **Residue left behind.** Water leaves nothing. Alcohol evaporates clean. Bleach leaves chloride and salt residue until rinsed. Detergent leaves film until rinsed thoroughly. Decontamination sprays vary by formulation and must be handled per instructions. A cleaning step that leaves its own residue simply trades one contaminant for another. **Throughput and automation.** For a single pipette at a bench, wipe-down methods are fast enough that throughput hardly matters. When the item being cleaned is a glass capillary used dozens of times per day, throughput decides the method, which is exactly the situation the ultrasonic method addressed. By enabling at least ten consecutive uses of a single patch pipette, that approach converted a discard-per-attempt workflow into a reuse workflow [3]. The parallel lesson for any lab is that cleaning method choice changes with usage volume, not just with contaminant type. **Safety and waste.** Water and dilute detergent are benign. Alcohol is flammable and needs ventilation and flammard storage discipline. Bleach is corrosive and should never be mixed with acids or ammonia. Choose the mildest solution that solves the actual problem, and you inherit the smallest waste stream. ## Worked Scenarios **Scenario 1. A shared cell culture room, end of day.** A group of six users shares four pipettes. The routine is a 70% ethanol wipe of the exterior after each session, a full detergent wash of the nose cone and ejector at the end of the week followed by distilled water rinse and dry, and filter tips used throughout. No bleach enters the picture unless a spill occurs. This allocation matches each solution to its contaminant, and it keeps the weekly detergent session short because residue never hardens. **Scenario 2. A trace-analysis method for bisphenols in milk.** A lab developing a QuEChERS and HPLC-QQQ-MS method for bisphenols A, S, F, B, and related compounds in condensed milk and breast milk faces analyte levels in the low nanograms per milliliter range in a very fatty, protein-rich matrix [1]. Carryover between samples is a first-order threat. The cleaning strategy is layered: dedicated pipettes reserved for standards versus extracts, filter tips changed between every sample, an immediate water rinse of anything reusable that contacts extract, and a standing rule that glassware and pipette contact parts get detergent washing and a final distilled water rinse before the next sequence. The QuEChERS sorbents such as Z-SEP Plus, PSA, C18, and EMR-Lipid clean the extract, but only pipette hygiene protects the extract from the pipette. **Scenario 3. An electrophysiology rig with high recording demand.** A lab running many patch attempts per day adopts ultrasonic tip cleaning within the recording bath, with detergent, so that each glass pipette serves at least ten consecutive recordings without harming slices or neighboring cells [3]. The detergent concentration and sonication cycle are validated by checking seal resistance and slice health, and the method pairs with automated patch-clamp to raise throughput. The right solution here is a process, not a bottle. **Scenario 4. A teaching lab pipette with suspected salt crust in the shaft.** An introductory lab finds one pipette dispensing erratically. The suspect is buffer wicking into the shaft after a student over-compressed the plunger. The response is immediate disassembly per the manual, inspection of the piston and seals, gentle cleaning of accessible metal and plastic internal parts with distilled water and, where the manual permits, dilute detergent, drying, fresh lubricant if the manufacturer specifies it, new seals if worn, reassembly, and a volume verification check before the pipette returns to service. The cleaning solution mattered less than the decision to open the instrument. ## Step-by-Step: Cleaning a Mechanical Air-Displacement Pipette ![Eight-step cleaning sequence from setup and exterior wipe through internal cleaning, drying, lubrication and a performance check.](/article-images/illustrations/best-cleaning-solution-for-pipettes-mechanical-cleaning-steps.webp) A mechanical air-displacement pipette draws liquid by moving a piston that compresses a fixed air column, so accuracy depends entirely on the piston, seal, and shaft staying clean, dry, and lubricated. The following sequence reflects standard practice documented in manufacturer manuals. 1. **Set up.** Work on a bench pad with the pipette set near its mid-range volume, wearing gloves, with a lint-free wipe, a small beaker of distilled water, dilute detergent, 70% ethanol, and the manufacturer's maintenance tools within reach. 2. **Exterior first.** Wipe the whole housing with 70% ethanol. Remove the tip ejector arm if your model allows it and clean behind it, because drips collect there. 3. **Nose cone and shaft opening.** Dampen a lint-free wipe with distilled water and clean the outside of the nose cone and the rim of the shaft opening. If residue is dried on, use dilute detergent, then rinse the area with a water-dampened wipe. Keep liquid out of the opening. 4. **Assess whether internal cleaning is needed.** Signs include visible liquid inside the shaft, stiff or gritty plunger action, volume drift on a quick check, or a known over-immersion event. If none apply, stop here and dry the exterior. 5. **Internal cleaning, only per the manual.** Disassemble exactly as the manual directs, typically by removing the tip ejector collar and unscrewing the shaft from the body. Withdraw the piston. Inspect the seal and O-rings. Clean the piston with distilled water on a lint-free wipe, and use dilute detergent only where the manual permits it. Never use alcohol on the piston or seal assembly unless the manual explicitly allows it, because alcohol strips lubricant. 6. **Dry completely.** Moisture inside the shaft causes corrosion and streaky delivery. Air dry or use a clean lint-free wipe on every component. 7. **Lubricate and reassemble.** Apply only the lubricant the manufacturer specifies, in the amount specified. Reassemble in reverse order and cycle the plunger several times through its full travel. 8. **Verify performance.** Run a quick gravimetric check if your lab has a calibrated balance, and flag the pipette for formal pipette calibration if the check looks off. A related decision is autoclaving. Many modern pipettes and their nose cones tolerate autoclaving at 121 degrees Celsius, but only when the manual lists the specific parts as autoclavable and only with the specified preparation, such as loosening the plunger or autoclaving disassembled parts in steam-permeable wrapping. Autoclaving an instrument that is not rated for it warps housings and cooks seals. ## Where Pipetting Technique Meets Cleaning Good pipetting technique reduces how often you need any of the solutions above, because most internal contamination enters through preventable errors. Keep the pipette vertical whenever it holds liquid, including while it sits in your hand between dispenses. Horizontal storage lets sample wick into the shaft, and the most common shaft contaminations in any teaching lab trace back to a pipette laid on its side with a loaded tip. Pre-rinse the tip when you need maximum accuracy with viscous or volatile liquids, which coats the inner wall and improves reproducibility, but recognize that pre-rinsing also means the tip, not the pipette, takes the residue. Change tips religiously rather than trying to decontaminate a used tip between samples. Aspirate slowly with viscous fluids, use reverse pipetting for foamy or volatile liquids, and never force the plunger past its first stop while liquid is loaded. Immersing the tip too deep, especially in foaming solutions, drags liquid up the outside of the tip and onto the nose cone, which is the entry route for most contamination. These habits are not accessories to cleaning, they are the front line. A lab with disciplined pipetting technique does a fraction of the cleaning a sloppy lab does, and its pipettes hold calibration longer. ## Pipette Calibration and Cleaning Cleaning and calibration are linked in both directions. A dirty pipette cannot be calibrated meaningfully, because residue on the piston or a sticking seal changes volume as the piston moves, so a calibration performed on a dirty instrument certifies a state that will not persist. Standard practice in most laboratories is therefore to clean, inspect, and, where applicable, relubricate before every calibration event, and to treat a failed calibration with unknown internal condition as a service event rather than an adjustment. Calibration itself works by dispensing defined volumes, most commonly water, and weighing them gravimetrically on a calibrated analytical balance, then converting mass to volume with the temperature-corrected density of water. The tolerances you test against, typically accuracy and precision limits for the nominal volume and reduced volumes such as ten percent of maximum, come from the manufacturer's specification sheet for your model and volume range. Multi-channel pipettes are checked per channel, because each channel has its own piston and seal. Cleaning extends the interval between calibrations in a very concrete way. Salts that dry on a piston abrade the seal with every stroke, and a scored seal leaks air, which shows up first as the pipette under-delivering at low volumes. Corrosion from bleach exposure produces the same failure. A pipette cleaned with compatible solutions and stored vertically is simply a pipette that stays inside specification longer. Most quality systems schedule calibration on a fixed interval and pull pipettes early when usage is heavy or history suggests drift, and your own maintenance log, including every cleaning event and any incident of liquid entering the shaft, is the record that justifies shortening or extending that interval. ## Common Mistakes and Limitations **Flooding alcohol into the shaft.** The single most common self-inflicted pipette injury in [molecular biology](/blog/careers/molecular-biology) labs is spraying the nose cone so enthusiastically that ethanol runs into the shaft opening and dissolves the piston lubricant. The result is a scratchy plunger and slow volume drift. Spray a wipe, not the pipette. **Assuming bleach is harmless because it is dilute.** A single 10% bleach wipe is fine. Weekly bleach wipes for a year corrode exposed metal and dull coatings. Bleach is a spot solution for biological risk, not a daily driver. **Using 100% alcohol for disinfection.** Higher concentration evaporates before it can act and is less effective against microorganisms than 70%. Buy 70% or dilute appropriately. **Rinsing detergent incompletely.** Detergent film transfers to the next tip and can inhibit enzymatic reactions downstream. If you see any suds on the second rinse, rinse again. **Reassembling wet or relubricating with the wrong grease.** Water left in the shaft corrodes, and petroleum jelly or generic grease is not the manufacturer's specified lubricant. Both mistakes show up as erratic delivery. **Autoclaving unverified parts.** Only autoclave components your manual lists as autoclavable, in the configuration the manual describes. **Confusing sample clean-up with instrument cleaning.** An analyst working through a QuEChERS method with sorbents such as Z-SEP, PSA, or EMR-Lipid [1], or running an aluminum oxide clean-up column packed into a Pasteur pipette [2], is purifying an extract. Those reagents have no role in cleaning the pipette itself, and labs sometimes inherit the vocabulary and assume otherwise. **Discarding when reuse was possible, or reusing without validation.** The patch-clamp field long defaulted to discarding glass pipettes after one attempt, and a detergent sonication method validated for at least ten consecutive uses with no harm to slices or other recorded neurons overturned that default [3]. The balanced position is to reuse where validation supports it and to discard where it does not. **Treating cleaning as a substitute for calibration.** Cleaning restores a maintained instrument. Calibration verifies it. Neither replaces the other. **Skipping the cleanliness check entirely.** The idea that pipette cleanliness deserves its own test is old enough to have its own literature. A test specifically designed to assess the cleanness of blood pipettes was published in a clinical laboratory journal as far back as 1961 [4], which tells you that generations of labs have recognized the need to verify, not assume, that a cleaned pipette is actually clean. A modern equivalent for a mechanical pipette is simple: after cleaning, dispense distilled water repeatedly and confirm free, smooth action, then return it to service or flag it for service. One general limitation applies to everything above. Individual instruments differ by manufacturer and model, and when your situation falls outside routine maintenance, such as suspected internal damage, repeated calibration failure, or contamination with hazardous biological or chemical agents, the right move is the manufacturer's service protocol or a qualified service provider rather than improvised chemistry. ## Frequently Asked Questions ### What is the best cleaning solution for pipettes overall? For most molecular biology work, the best overall routine is distilled water for immediate rinses, 70% ethanol for exterior disinfection, and dilute detergent with a thorough water rinse for scheduled deep cleaning, escalating to 10% bleach only for biological decontamination. The best single solution does not exist because residue removal, disinfection, and decontamination are different chemistry. ### Can I use regular dish soap to clean pipettes? Plain, unfragranced dish soap in a dilute solution can work in a pinch, but laboratory-grade detergent is the safer default. Many household soaps contain lotions, fragrances, or abrasives that leave films or scratch housings, and any soap must be rinsed completely with distilled water. ### How often should pipettes be cleaned? Wipe the exterior with 70% ethanol at the end of each work session, perform a detergent wash and water rinse on a weekly or monthly schedule depending on usage, and clean internally immediately whenever liquid enters the shaft or performance changes. Heavy-use and shared instruments justify shorter intervals. ### Is bleach safe on pipettes? Dilute 10% bleach is safe for occasional exterior decontamination of hard plastic and glass surfaces, followed by a distilled water rinse and dry wipe. It is not safe for repeated contact with metal pistons or internal parts, and it should never be run through the shaft. ### Can ultrasonic cleaning damage a pipette? Sonicating a fully assembled mechanical pipette drives liquid past the seals and destroys lubrication and calibration, so do not do it. Ultrasonic detergent baths are appropriate for glass capillaries and sample-contact glassware, where a validated sonication method lets patch-clamp pipettes be reused at least ten times without harming the preparation. ### Does cleaning replace pipette calibration? No. Cleaning keeps the piston, seals, and shaft in the condition that allows accurate delivery, while calibration gravimetrically verifies that the delivered volumes still meet the manufacturer's specification. Both belong in the same maintenance program, with cleaning preceding every calibration event. ## Sources 1. [Method Development for Selected Bisphenols Analysis in Sweetened Condensed Milk from a Can and Breast Milk Samples by HPLC-DAD and HPLC-QqQ-MS: Comparison of Sorbents (Z-SEP, Z-SEP Plus, PSA, C18, Chitin and EMR-Lipid) for Clean-Up of QuEChERS Extract.](https://pubmed.ncbi.nlm.nih.gov/31159388/) 2. [Comparison of extraction techniques of robenidine from poultry feed samples.](https://pubmed.ncbi.nlm.nih.gov/19073106/) 3. [Immediate reuse of patch-clamp pipettes after ultrasonic cleaning.](https://pubmed.ncbi.nlm.nih.gov/38238544/) 4. [A solution for testing the cleanness of blood pipettes.](https://pubmed.ncbi.nlm.nih.gov/14463768/)