A few months ago, I got an email from a young researcher at a small biotech lab. She’d been working on a critical stem cell line for six months—her whole project depended on it. Then, during a routine passage, her cell culture samples got contaminated. When she checked her pipettes, she found tiny residual crumbles of dried medium stuck inside the tips, left over from a previous use with a different cell line. That mistake cost her three weeks of work, and she told me it almost made her abandon the project. Cell Culture Consumables
As a cell culture consumables supplier who’s spent 12 years in this space, that story sticks with me. I see this kind of thing all the time, not because researchers are careless, but because pipette sterility feels like a basic step—until it’s not. Pipettes are the unsung workhorses of cell culture; they transfer every drop of medium, reagent, and cell suspension that keeps your cultures alive and experimental results reliable. Even a single non-sterile pipette can introduce bacteria, fungi, or cross-contaminated cell lines that derail months of work. So let’s talk about how to get this right, not just with generic tips, but with the practical, science-backed steps that our own customers rely on every day.
First, let’s clear up a common misconception: “sterile” isn’t just a label on a box. For disposable pipettes, which are the most commonly used in routine cell culture, sterility means they’ve undergone a validated sterilization process—usually gamma irradiation, which is the gold standard for plastic lab consumables— to eliminate all viable microorganisms, including spores. But here’s the catch: that sterility only holds if the packaging remains intact until you use it. The mistake many researchers make is pulling a pipette out of its wrapper early, leaving it out on the bench while they prepare samples, or even reusing pipettes that were meant to be single-use (I’ve seen this too, from grad students trying to stretch a budget on a tight supply). Even if you sterilize a pipette via autoclave, which we don’t recommend for standard disposable pipettes because it can warp the plastic and create tiny crevices that trap contaminants, you have to do it correctly. Autoclaving parameters—temperature, pressure, time—vary based on the pipette material, and if you mess that up, you’re not actually achieving sterility.
Let’s start with the basics that most labs overlook when handling sterile pipettes: storage and prep before use. When your shipment of sterile pipettes arrives, don’t just dump them in a cabinet under the bench. I recommend storing them in a cool, dry, dark place—ultraviolet (UV) light from overhead lights or lab windows can break down the gamma-irradiated sterility barrier over time, especially if the packaging is damaged. Keep them in their original, sealed packaging until you’re ready to open them, and never move the package around while it’s open. That’s how dust, airborne microbes from the lab’s air handler, or even lint from your lab coat gets inside the wrapper and onto the pipette tips.
Before you even step into the laminar flow hood, there are a few non-negotiable steps. First, sanitize your hands with 70% ethanol, and let them dry completely—wet hands can leave moisture that transfers to the pipette packaging. Then, spray the outside of the pipette wrapper with 70% ethanol, wipe it down with a lint-free Kimwipe, and let it sit for 10 seconds before opening. This kills any contaminants on the outside of the package that could fall into your hood when you peel it open. While you’re at it, make sure your laminar flow hood has been running for at least 15 minutes before you start—fresh air from the hallway or a door opening can blow contaminants into the hood, even if it’s working correctly. And for the love of all things cell culture, don’t leave the hood sash open wider than necessary; that disrupts the airflow and lets in outside microbes.
Now, opening the pipette correctly is a make-or-break step. I’ve watched new grad students tear open a wrapper like they’re opening a candy bar, grabbing the pipette tip end with their fingers before it’s even fully out. No—you want to open the wrapper on a flat surface inside the hood, ideally about 6 inches from the front edge of the hood’s work surface (that’s where the airflow is cleanest). Most sterile pipette wrappers have a tear notch that opens to two flaps: one that holds the pipette, and one that’s designed to stay closed. Peel one flap back slowly, keeping the pipette inside the wrapper until you’re ready to take it. When you grab the pipette, only touch the end where you insert it into the pipette controller. The tip end—where you attach the pipette tip—should never touch anything: not your gloves, the hood surface, a tube rack, or even another pipette. That’s how cross-contamination happens. If you accidentally touch the tip, toss that pipette and grab another. It’s not worth the risk.
Speaking of pipette tips—wait, did you know that even sterile pipettes can have non-sterile tips if you use the wrong kind? A lot of labs use universal pipette tips, but if you’re working with sensitive cell lines that can be contaminated by even trace microbes, you need filter tips. Filter tips have a hydrophobic filter at the top that prevents aerosols from getting sucked up into the pipette shaft, and also blocks any contaminants from the pipette itself from touching your sample. I’ve had a researcher tell me they thought filter tips were a “luxury” until they lost a whole transfection experiment to an airborne fungal spore that got into their sample through a non-filter tip. Now, for routine work, filter tips are non-negotiable in my book. And always make sure the tips are packaged separately, not loose in a box—loose tips can get contaminated just from being jostled around in transit or storage.
Once you’ve attached the tip to your pipette, how you pipette also affects sterility. You might think it’s just squeezing the plunger, but there’s a right way to do it for cell culture. When aspirating sample, don’t dip the tip deeper than 1 cm into the liquid, and never let the tip touch the side of the tube (unless it’s a deep well plate, but even then, be careful). If you touch the tube wall, you’re picking up any contaminants that might have settled there. Also, never eject the liquid from the tip by force—slowly release the plunger so you don’t create aerosols. Aerosols are one of the biggest culprits for cross-contamination in cell culture; they can travel several feet through the hood and land on another sample, or even on your pipette, and you won’t see them. I’ve had samples get contaminated three days after a pipette use because an aerosol was left on a tip I’d put back on the bench (never, ever put used pipettes or tips back into the sterile packaging—throw them away immediately in a designated pipette waste container inside the hood).
Another thing people don’t talk about: reusing pipettes. We sell both disposable and autoclavable pipettes, but I always advise our customers to use disposable sterile pipettes for routine cell culture. Even if you autoclave a pipette, you can never be 100% sure you’ve killed all contaminants—autoclaving relies on consistent temperature and pressure, and if the pipette has any scratches or dents, it’s impossible to get all the crevices. Plus, autoclaving can damage the pipette’s calibration, so your volumes will be off, which is another problem for cell culture. If you’re working on a project that requires large volumes, our disposable pipettes are designed to be affordable enough that you don’t have to reuse them, and they’re already pre-sterilized so you don’t have to worry about the autoclaving step.
What about if you’re working with multiple cell lines in the same hood? That’s when cross-contamination is a real risk, and pipette sterility is even more critical. The rule here is: use a new sterile pipette for each cell line. Don’t even rinse a pipette between lines—you’ll leave residual cells or medium that can contaminate the next culture. And always process the most sensitive or most valuable cell line first, before moving to less important ones. That way, if something does get contaminated, you don’t lose your entire valuable culture.
I also can’t talk about pipette sterility without mentioning quality control. As a supplier, we test every batch of our sterile pipettes to make sure the sterility is valid—we do microbial challenge tests to ensure that gamma irradiation has killed all test organisms, and we check packaging integrity to make sure it doesn’t have any holes that would let contaminants in. But that doesn’t mean you don’t have a role in checking too. When you open a new box of pipettes, do a quick visual check of the packaging. If it’s torn, dented, or has any moisture inside, don’t use those pipettes—send them back. Moisture inside the packaging is a sign that the sterility barrier has been compromised, and that means contaminants could have gotten inside. I’ve had customers send back whole pallets of pipettes because a shipping box got rained on, and the packaging was damp. It’s better to be safe than sorry—one contaminated experiment is way more expensive than a few extra pipettes.
Let’s circle back to that young researcher who emailed me a few months ago. After she lost her stem cell line, she called us, and we walked her through a step-by-step check of her whole pipette workflow. Turns out, she was opening her pipettes before she entered the hood, leaving them out on the bench while she prepared her medium, and reusing pipettes for two different feeder cell lines. She adjusted all those steps, switched to filter tips, and three months later, she emailed me saying her stem cell line had been stable for two passages, and she’d got her first set of reliable data. That’s what this is all about—making sure your hard work doesn’t get derailed by a preventable mistake.
At the end of the day, pipette sterility isn’t a complicated science, but it’s a consistent process that needs to be part of every cell culture routine. It starts with using pre-sterilized, high-quality pipettes from a supplier you can trust, handling them correctly from packaging to use, and never cutting corners when it comes to avoiding cross-contamination. If you’re struggling with consistent sterility in your lab, or if you need guidance on the right consumables for your specific cell culture work, our team is here to help. We understand that every cell line is different, and we can walk through your workflow with you to find solutions that fit your needs and your budget.
Medical Consumable References
- Freshney RI. Culture of Animal Cells: A Manual of Basic Technique and Specialized Applications. 7th ed. Wiley-Liss; 2016.
- Caputo C, Atzori L, Cossu A, et al. Contamination control in cell culture: a practical review. J Biol Methods. 2017;4(2):e78.
- ISO 11135:2014. Sterilization of health care products—Requirements for validation and routine control of ethylene oxide sterilization. International Organization for Standardization; 2014.
- American Type Culture Collection (ATCC). Cell Culture Basics: Contamination Prevention and Detection. ATCC Technical Bulletin; 2020.
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