First, we must understand that in our planning we want to leave the underlying scientific process of our experiments untouched.

In other words, the goal is to drive change where it doesn’t affect our science.

This is why sustainability-related change doesn’t mean significant investments of additional time, effort, or risk.

The biggest reason we are afraid of change is that we prefer to keep things steady because we are already busy with everything else.

Understanding Perceived Risk

We rarely learn how to optimize, how to drive change, and sometimes we don’t even know why our protocols work.

This makes change feel overwhelming. It makes our mind rationalize irrational concerns.

Generally, it is often possible to find a technical solution – the difficult part is clearly expressing the kind of anxiety that holds us back.

To allow us to think accurately, we need to identify and label the anxiety that we are often not aware of.

> Otherwise, we see dangers everywhere or nowhere. But we want to see the actual risks and address them.

Let us therefore look at some examples.

• A common perceived risk is overlooking a danger and thereby causing contamination. However, if we plan our optimization ahead and review it on two different days, it is very unlikely that we would make such a blunder.
• Secondly, one might be afraid of being distracted and thereby making mistakes. For example, when reusing the same pipette tip and causing contamination. While this can always happen (why else would cell cultures get contaminated), one might even argue that because we work more sustainably, we pay more attention to best practices and focus more on our experiments.

• Also, some fear that after a change the experiment no longer works as it did previously. But if this were the case, we can simply return to the original method. Of note, investigating surprising failures has been the starting point for ​breakthrough​ findings.

The True Danger

This means most dangers introduced to scientific processes are linked to human failure, not the riskiness of optimization.

In other words, we (rightfully) perceive excessive and therefore blinding eagerness to save resources as the danger.

However, this eagerness fades once we understand it’s not about reaching a zero footprint.

Many scientists fear that sustainability will fundamentally change their science because they assume we must radically cut resources. This is not the case. The goal is optimization.

This is also why radical changes are often not the most sustainable – for example, switching all plasticware to glass is not necessarily efficient, because glass has a large footprint itself, sometimes greater than reused plastics.

My goal is to make you see sustainability-driven changes as a synonym for optimization.

We essentially optimize; we don’t change. And that also means it’s not primarily about the environment – it is about optimizing workflows.

The point is that saving time, chemicals, or plastics naturally translates into sustainability.

The Levels We Operate On

This means most dangers introduced to scientific processes are linked to human failure, not the riskiness of optimization.

Unless we choose to use innovative technology (e.g., switching from conventional assays to SPR), the underlying methodology always stays the same.

That means we might, for example, change the items that we use—this is a change in procurement. We might purchase bio-based materials that are of the same quality.

We may change how we handle our items. For instance, instead of using a 50 mL tube, we use a 15 mL tube. We may also change how we handle our instruments—for example, turning them off during the weekend or optimizing their settings, such as when we use a microscope.

Finally, we can choose to optimize our experimental design—such as when working with mice—so that in the end we have a different setup, for example by changing sample numbers or investigation schedules.

However, none of these require us to change fundamentals of our protocols or rely on assumptions about biological or chemical processes we cannot be certain about.

In fact, protocol optimization, when done as a team, doesn’t require senior expertise from every member. A bachelor’s student can make a suggestion that is valued by a postdoc, as I have experienced myself. In other words, experience should validate proposed changes, but optimization simply requires a thorough understanding of the underlying science.

How It Benefits Science

In essence, we can reframe the question of sustainability and safety.

Sustainability efforts should remove unnecessary steps—steps that are often inherited from outdated purposes because protocols were not optimized or were reused for something else.

This can have significant effects because, in many cases, we have not tapped into the potential for optimization at all.

In many academic settings, protocols were never optimized, and thus adjustments can make huge differences: reductions of 30–50% in solvent use, 60–80% in plastic waste, and sometimes cutting analysis time in half.

Even in industry, where protocols tend to be more optimized, these optimizations usually focus on the scientific aspects, not the handling of items – an area that represents a major opportunity for cost savings.

My goal is to make you see sustainability-driven changes as a synonym for optimization. We essentially optimize; we don’t change.

And that also means it’s not primarily about the environment—it is about optimizing workflows. Saving time, chemicals, or plastics naturally translates into sustainability.

Once we adopt this perspective, we also see the scientific advantages.

We often assume that our experiments measure our target processes without noticing how unnecessary manipulations alter the system.

Sustainability removes unnecessary steps—such as diluting solutions in intermediate tubes—thereby reducing opportunities for contamination or error. In other words, reducing unnecessary steps improves scientific quality.

How To Do It

To implement change safely, following five core principles might help:

• Differentiation There is no single strategy that works everywhere. Reusing a pipette tip might be appropriate for certain controls where only the concentration differs but the analyte is identical, whereas it would not be appropriate for specific sample types. We should avoid oversimplification.

• Stepwise implementation Many protocols offer multiple points for optimization. Although it may be tempting, the best approach is stepwise change. This reduces cognitive burden and ensures that if unexpected difficulties arise, we can trace them back and handle them on the spot.

• Mindset When implementing change for the first time, we need the right mindset. This means planning and analyzing the change beforehand so that we can remain fully present when conducting the experiment. It also means working in a good flow—not being distracted by worries about colleagues’ reactions or anxiety about the change itself. Confidence, focus, and concentration are essential.

• Experience We must be sufficiently familiar with the protocol before optimizing it. Protocols that are handed down should first be learned and implemented as they are. Then, optimizing includes talking to lab colleagues about potential difficulties and thoroughly reviewing literature to see whether similar changes have been reported.

• Controls This involves performing trial runs to verify whether optimizations are still valid. Then, it’s about documenting changes through controls – in cell culture, for example, this means checking whether cells grow with the same morphology, speed, and metabolism when a new dish type or dish-reuse strategy is employed.

Applying The Knowledge

The key is not to change a running system in the middle of an experiment.

Instead, change should be implemented after an experimental series is completed or when a new project is started.

Most often, optimization is missing due to psychological barriers.

One such barrier is insufficient trust in oneself.

A very different one is the reluctance to accept that improvements already exist, as this would mean admitting that one could have been more efficient for a long time or that someone else might find a solution one did not.

When leaders raise doubts, resistance might stems from distrust in a person, not distrust in the change itself (although expressed as such).

However, if you as a supervisor doubt a person’s ability to optimize, you might ask yourself whether you truly trust that person to conduct experiments properly – and if not, why. Then, of course, it is your responsibility to grow them or remove them. Otherwise, what does this imply for the future of the project or the group

Nevertheless, if doubts remain after meticulous planning, change should be aborted. This may indicate that the optimization has the wrong target, impacting aspects of the underlying process.

It might also suggest that the protocol itself is so fragile that it yields essentially artificial data, which requires you to make a call: fundamentally rework the protocol or live with it.

As scientists, we constantly work on new projects and approaches, meaning we are inherently used to change. However, the circumstances matter and this is what we need to express.

If you want to convince yourself, colleagues, or supervisors:

1. Adhere to best practices: research the literature, plan the change carefully, and run a trial.

2. Assure superiors that you will invest the extra time required to avoid losses in productivity, even though long-term benefits are almost always observed.

3. Prove that you are capable of managing the change by clearly articulating what we know and plan to do, even if it feels obvious or trivial because others don’t know what you do.

Once again, the biggest challenge is not the science. it’s the psychological barriers we are rarely aware of.

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References

Penndorf, P., et al., 2023. A new approach to making scientific research more efficient – rethinking sustainability. FEBS Letters, 597(19), pp.2371–2374. doi:10.1002/1873-3468.14736.

Farley, M., et al., 2023. Re-use of laboratory utensils reduces CO₂ equivalent footprint and running costs. PLOS ONE, 18(4), e0283697. doi:10.1371/journal.pone.0283697.

Cushman, F., et al., 2020. Rationalization is rational. Behavioral and Brain Sciences, 43, e28. doi:10.1017/S0140525X19001730.

Alves, J., et al., 2020. A case report: insights into reducing plastic waste in a microbiology laboratory. Access Microbiology, 3(3), 000173. doi:10.1099/acmi.0.000173.

Kilcoyne, J., et al., 2022. Reducing environmental impacts of marine biotoxin monitoring: A laboratory report. PLOS Sustainability and Transformation, 1(3), e0000001 doi:10.1371/journal.pstr.0000001.

Mazzali, D., et al., 2025. Sustainable and surfactant-free synthesis of negatively charged acrylamide nanogels for biomedical applications. Macromolecules, 58(3), pp.1206–1213. doi:10.1021/acs.macromol.4c02128.

From the very beginning, I was one of the few who openly talked about misconduct.

Ever since the start of my career in sustainability, I have worked with different companies.

This is how I fund our activities, and of course, this is also how I gain access to information that nobody outside would normally see – information that I am then able to share with you.

But we also know this is why many people deviate from their original path of boldly addressing shortcomings.

Why? Because at some point, money becomes too important.

Since this has not happened to me, and as I am one of the few people in the field with the necessary technical expertise, I want to talk about something I often address in my advisory practice.

I want to give you some examples because I want to help you avoid falling prey to this kind of misguidance.

Pitfall #1: Negative Carbon & “Certifications”

A major trend at the moment is materials made from biogenic sources – for example, plant waste streams.

But one of the biggest problems I see is the claim of a negative carbon footprints.

How is that possible? I would argue it isn’t.

It’s just a mathematical trick.

Yet, combined with clever marketing it becomes misleading as we see only the final number, not how it was calculated.

Here’s how it works: biogenic carbon refers to carbon that plants absorb from the atmosphere and store in their biomass.

I.e., if we make materials from plants, we’re storing atmospheric carbon and therefore ending up with a negative footprint.

However, this thinking only applies when materials are used for a long time (like in construction for 50-150 years).

But there is the workaround: companies are allowed to conduct only cradle-to-gate life cycle analyses, i.e., from material sourcing to the point the product leaves the factory.

That allows them to claim biogenic carbon storage without accounting for what happens after use – even if the item is incinerated just two days later.

Even worse though is that companies can certify these numbers.

For instance, ISCC certification is available for such claims.

When we hear “certification,” we assume reliability. In my view, what ISCC is doing here is either naïve or negligent.

> They allow companies to certify cradle-to-gate analyses, enabling misleading marketing claims.

And to make matters worse, companies can back these claims with ISO certifications few consumers are educated about.

The issue is twofold: many ISO standards merely tell companies what to do, not how to do it. They outline for example that “Cradle to …” boundaries have to be defined but not which ones.

On top of that, not all ISO standards are certifiable. Some are just frameworks, especially those related to life cycle assessments.

That means: in practice, companies can design their assessments to get the most favorable results, calculate within that limited setup, and still get certified.

Especially in this case, certification only verifies their bookkeeping, not the applicability of their sustainability claims.

Fittingly, ISCC has already faced scrutiny:

And even if we conduct a full life cycle assessment:

We can refer to the “neutrality assumption” (meaning the carbon fixed by plants equals the carbon released, setting emissions from those to 0).

And still, we have emissions from manufacturing, transport, and end-of-life treatment. Every product has a footprint.

A “negative” footprint implies that the more you buy, the better for the planet which is simply not true.

PS: Apart maybe from a few selected waste streams that end up in the construction sector.

Pitfall #2: False Sense of Transparency and Information

I generally support take-back programs. This is also why I’ve promoted them previously.

However, a line is crossed when marketing goes too far, because none have convincingly proven their sustainability officially.

While a fraction of companies at least share where their recycling plants are located, they rarely disclose transport details.

If small batches travel long distances, the transport footprint can outweigh the recycling benefit.

For the larger fraction that doesn’t disclose their recycling sites, it’s even worse.

Since waste management chains are often opaque, there’s always the risk that your waste is incinerated or landfilled instead of recycled.

I suspect that most companies lack data to verify where their collected waste actually ends up.

In short, we’re working with a black box – nobody really knows the true footprint of these programs.

As if that weren’t enough, some of these programs claim “closed-loop” recycling.

In reality, they use a fancy term and show you a facility – but not its limitations.

On average, recyclable plastics can only be recycled about five to eight times before their quality degrades too much.

Chemical recycling could, in theory, extend this – but it’s still not scalable and has a much higher footprint.

So “closed-loop” usually means a single loop before virgin materials have to be added – not an infinite one.

Pitfall #3: Best-Case Assumptions

Companies with great innovations often make sustainability claims that are theoretically correct but practically unrealistic.

They assume the best-case scenario for their product, often based on very local or purely theoretical circumstances.

Moreover, there’s no regulation forcing them to compare against a defined baseline.

That means they can select any reference point that makes their results look good – claiming large “savings” that may not hold up in reality.

For example, when a product is labeled “biodegradable,” we rarely know what that means in practice.

Definitions vary, composting facilities are scarce, and contaminated lab waste can’t be composted anyway.

Manufacturers assume ideal conditions, but in real life, these products are often incinerated or landfilled.

If they assume landfilling, they’re allowed to compare against a product that is incinerated and whether they consider “waste-to-energy” or simple burning is up to them.

The same applies to manufacturing claims about greener plastics.

PLA and other new materials come with additional environmental burdens but whether and how these are considered is up to the company.

Applying The Knowledge

Sometimes, working with a sustainability advisor can be crucial.

Why? Because following misleading marketing claims might lead you to choose a product that sounds greener but isn’t.

In the future, you might have to report your sustainability practices to a funding body or agency.

Using flawed data could mean your proposal is rejected or sent back for correction, creating a huge amount of extra work that you have to handle.

Therefore, pay attention:

• “Verified by …” ≠ objective truth. Certifications are helpful, but they don’t guarantee objectivity.
• What don’t you know about the process you invest in?
• What is the best-case scenario vs. what really happens? Follow impressive numbers – but ask what baseline was used?

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Written by Patrick Penndorf

Join our Educational Series | Connect with me through LinkedIn

This guide will explain to you the impact of freezers, how you can make them greener and how to maintain them properly.

Concise, applied and easy to understand.

Let’s start with the fundamentals: the impact of fridges and freezers:

To this day, there is not a single comprehensive life cycle analysis available for a -80°C ultra-low temperature (ULT) freezer. The same goes for lab fridges. ​

​Therefore, I went through the available data and, where necessary, translated findings from commercial fridges to our situation. Although tedious, the data was surprising:

​​Manufacturing Impact

​A typical household fridge weighs 60–150 kg and is made from a wide range of materials, including stainless steel, aluminum, copper, polystyrene, tempered glass, acrylics, and polyurethane foam.

​The carbon footprint of manufacturing such a fridge is around 200–800 kg CO₂e. Walk-in refrigeration units or those used in supermarkets have accordingly higher impacts.

​Lab freezers, are somewhat heavier: a 500 L -80°C freezer weighs 240–340 kg, meaning its estimated manufacturing footprint is 600–900 kg CO₂e.

​Interestingly, larger 700+ L models weigh roughly the same (260–315 kg), resulting in a similar manufacturing footprint.

​Energy Consumption – The Biggest Factor

Most of a freezer’s footprint comes not from its production but from its electricity use. The ongoing impact is as follows:

​Household fridge:

• 1–2 kWh/day → 0.64 kg CO₂/day
• 77–234 kg CO₂/year

-80°C freezers (500+L, assuming 341 g CO₂/kWh in Germany):

• Very old model: 36 kWh/day → 4,480 kg CO₂/year
• Newer model: 12 kWh/day → 1,493 kg CO₂/year
• Best model available: 7 kWh/day → 871 kg CO₂/year

​Of note, these are compressor-based fridges—i.e., they use a compressor for cooling. For -70/-80°C, these are the most common models. However, some units use liquid nitrogen (especially for -130°C) for cooling, resulting in a very different life cycle composition.

​Refrigerants

In short, a refrigerant is a substance used in cooling systems to absorb and release heat, enabling refrigeration and air conditioning.

It cycles through phase changes (liquid ↔ gas) to transfer heat efficiently.

Why does this matter? Because refrigerants have an enormous climate impact when leaked. For instance, assuming a 50% refrigerant recovery during disposal:

• A small -20°C freezer (R134a) → 71.5–143 kg CO₂e impact at disposal
• A -80°C freezer (R404 & R508) → 2,000–3,631 kg CO₂e impact at disposal

Due to a 2–15% leakage rate per year, full recovery is impossible, and occasional refilling is required. On top of that, illegal disposal and improper handling remain widespread.

​End-of-Life

The end-of-life impact, including disposal, plays a smaller role in the overall footprint.
Estimates range between 0.25–15 kg CO₂e.
(Deeper discussion about potential false assumptions in our Slack)

Transportation to the landfill is likely a significant contributor. Additionally, we need to account for the energy consumption of hulk shredding (approx. 144 J/kg) and material separation.

​Typical recycling rates look something like steel, aluminum, and copper: Recycled at 37%, 32%, and 22% by weight, respectively. Plastics and residue: 20% incinerated, 80% landfilled.

When Should You Exchange Your Freezer?

A fridge has a total footprint of about 2000-8000 kg CO₂e over a 10-year lifespan. For a -80°C freezer, that number is closer to 20 000–
50 000 kg CO₂e. (More in our free Slack).

Let’s summarize:

• Financially: The cost of a new freezer amortizes over 5–10 years, depending on energy prices.
• Environmentally: The footprint amortizes in 2–5 years due to massive energy savings.

Compared to a 500+L model, the 700+ L energy consumption increases by roughly 30%, meaning faster amortization in environmental terms.

​Bottom line:

• If your freezer is 15+ years old → Definitely replace it.
• If your freezer is 10+ years old → Replacement is likely a good decision.
• If your freezer is 5+ years old → Debatable, analysis recommended

Making Freezers More Sustainable

Quick Aside: The global freezer market in 2022 was estimated at a staggering $4.7 billion. However, by 2030, estimates predict it will grow to between $7.4 billion and $12.7 billion. While the EU and US markets are the largest, Asian markets are expanding rapidly. Nevertheless, it is notable that Africa and South America—despite having significantly hotter climates—accounted for less than 5% of the market in 2024

As we discussed, the biggest environmental impact of ultra-low temperature (ULT) freezers comes from their use phase. ​ But how big is their energy consumption compared to other lab equipment? ​

​The average energy consumption of a freezer per year ranges between 2.5 – 9.1 million Wh. An average American household consumes between 9 and 11 million Wh per year. Thus, only the older models consume more energy than an entire home!

​Now that we have a sense of scale, let’s explore how to minimize freezer energy consumption as much as possible.

​Purchasing

​At first, choosing a freezer with lower energy consumption will be environmentally beneficial.

​Modern ULT freezers have become significantly more energy efficient over the past few decades.

A 500+ L model from before 2000 can be expected to use up to 36 kWh/day, whereas models from the 2000s–2005s typically consume around 19 kWh/day.

​More recent models have further improved efficiency, consuming between 7–12 kWh/day, with the most efficient freezer currently available (PHCBI) using just 4.99 kWh/day.

​These efficiency gains are the result of multiple technological advances:

• Improved heat exchange designs optimize cooling performance.

• Vacuum-insulated panels have replaced polyurethane foam, reducing heat loss.
• Variable-speed and dual-stage compressors adjust power dynamically, operating only at the minimum required level rather than running at full power constantly.

However, even the best freezers age. Every year, energy consumption increases by approximately 1-3%, which can easily translate to an additional 8.75 kg of CO₂e per month.

​​​Adjusting Set Temperatures

The single most impactful change a lab can make is adjusting freezer temperatures from -80°C to -70°C or lower.

Based on studies from the University of Copenhagen, together with an investigation by Farley et al., and manufacturer data, we can confidently say that raising the temperature from -80°C to -70°C reduces energy consumption by 22–29%!

​

​In fact, -70°C was the standard for decades before improvements in cooling technology made -80°C the norm—not for scientific reasons, but because it became a marketing advantage.

​Published studies further confirm that sample stability is not affected:

• ​​Espinel-Ingroff et al. successfully recovered 6,000+ yeast and 300+ mold samples after 10 years of storage at both -70°C and -80°C.

• ​Landor et al. found no significant degradation in DNA and RNA stored at -70°C, with only minor 260/230 ratio variations, all well below standard deviation ranges.
• Paraoxonase-1 enzyme activity remained unchanged after one year of storage at -70°C

​Even manufacturers acknowledge this—QIAGEN officially recommends storing RNA at -70°C.

​And it’s not just ULT freezers—adjusting standard freezers from -25°C to -20°C has been shown to reduce energy consumption by 20%, as reported by EPFL in Switzerland.

​The Role of Proper Setup and Maintenance

​Aside from temperature adjustments, proper installation and maintenance can significantly reduce a freezer’s energy consumption.

​A study by Gumapas et al. analyzed four freezers (20–25 kWh/day) under different conditions and found several key factors:

​Controlling Ambient Temperature

• Each 1°C increase in room temperature leads to 18 kWh of extra energy use per month, releasing an additional 9.27 kg of CO₂e. (+ don’t place freezers at windows/in the sun)

​Avoiding Dust Accumulation

• Regular cleaning can save 211 kWh per month, preventing 108 kg of CO₂e emissions.

• ​Declogging filters alone reduced energy consumption by 117 kWh per month, or 60 kg of CO₂e.

Ensuring Ventilation

• In their case, poor ventilation increases the duty cycle by 4%, leading to an additional 85 kWh of electricity used per month (equivalent to 51 kg of CO₂e).

​Removing Ice

• Heavy ice accumulation can increase energy use by up to 50%, forcing the compressor to work harder to maintain low temperatures.
  (Next week more on how to clean your freezer)

​​Some Important Questions:

​Should You Purchase Larger Freezers?

​Older studies suggest that larger freezers were more about 13% for efficient. However, this depends on the manufacturer. While such a difference is still exists for PHCBI models, they do not in Eppendorf ULTs.

​Of note, upright freezers tend to consume less electricity than chest freezers.

​​How to Convince Colleagues to Switch to -70°C

​If your lab is hesitant, the best strategy is to start with one freezer and demonstrate that it is completely safe.

​In fact, due to reduced compressor load, freezers set to -70°C are less likely to fail under suboptimal maintenance or when air conditioning struggles to keep up.

​If you would like to demonstrate how much money can be saved though purchasing more efficient freezers, PHCBI has a simple online tool (just note that their competitors have newer models with smaller energy consumption).

​Maintaning Freezers Sustainably

​If your freezer fails, you have only a few hours before it reaches temperatures that could harm your samples. Therefore, prevention is key.

Although Proper freezer and sample maintenance can save a lot of energy, it also will:

A) Safeguard samples from being lost.

B) Prevent unnecessary failures and reduce search time.

C) Maintain freezer health, as frequent openings and insufficient cleaning are the main causes of failure.

​Sample Organization

​Each time the door is opened, warm air enters, forcing the compressor restore lower temperatures. Additionally, this leads to ice buildup on the lid, increasing the risk of compressor overload due to constant warm air exposure.

​However, a well-organized system comes with further advantages for your science:

​1. Reduces time needed for searching samples.

​2. Prevents samples from being lost or forgotten (consider the high turnover of people in laboratories).

​3. Ensures detailed sample metadata, which, if missing, might lead to misleading results:

• Experiment-specific notes (e.g., genetic variations in bacterial strains, passage number for cell cultures, source of biological material).
• Treatment conditions (e.g., exposure to chemicals, storage buffer composition).
• Tracking expiration dates to prevent unnecessary rework or contamination from degraded samples.

​How to?

​The simplest and cheapest approach is an Excel-based tracking system.

​Here is our short guide on how to create such a system, along with a ready-made template.

​In short, you use one sheet to track each type of sample/reagent and one for each freezer.

​While an Excel system is highly customizable and cost-effective, it requires manual updates and is prone to human error.

​Otherwise, you can decide for an automated solution – using a sample management software and QR codes that are readable with with a smartphone.

​These systems also enable enhanced data storage, including full experimental history and direct links to references or protocols.

​Some software options even synchronize with electronic lab notebooks (ELNs) or provide real-time tracking of reagent usage

​However, no matter how well-organized your freezer is, you will still have to open it. Each time you do, humid air enters.

​​How to Maintain & Clean Your Freezer

​The most common cause of freezer failure is compressor overload, often due to blocked ventilation or insufficient heat transfer.

​While the former is typically caused by dust accumulation on the filter, the latter is linked to dust on the condenser coils. Additionally, ice buildup on the gasket/rubber seal can prevent proper closure, forcing the freezer to run constantly to compensate for the air exchange.

​The key steps to keeping your freezer in good condition are:

• Cleaning the condenser filter (usually located at the base/back).
• Cleaning the condenser coils.
• Defrosting the freezer.
• Cleaning the vacuum relief port/auto vent port.
• Discarding unnecessary samples

​

Here’s our easy and concise guide on how to clean your ULT freezer effectively.

​Also, this NIH video explains quite well how to clean the filters.

​Some Tips For You

​While these activities do not generate new data, they ensure your work runs efficiently and that samples remain well-preserved.

​Even if your lab members aren’t actively participating, I highly recommend setting up a sample organization system and basic freezer maintenance procedures for yourself. The benefits will become clear once you’ve worked on a project for more than six months.

​If you receive positive feedback from your team, use that momentum:

• ​Schedule routine cleaning in your lab’s maintenance calendar.

• Ensure guides on freezer cleaning and sample documentation are easily accessible. Ideally, store them in the same folder as the actual software/Excel file.
• Provide proper training for team members. If people feel uncertain, they might either avoid the task or do it half-heartedly, leading to mistakes.

​Finally, consider joining initiatives like the International Freezer Challenge.

Bonus

As many labs face challenges defrosting their freezers due to limited capacity for temporarily storing their samples, I’ve found a company that offers an excellent solution.

They also provide secure sample storage at various temperatures, with full certification.

Read more here: https://readvance.kit.com/posts/sample-safety-transport

Some clear-eyed orientations to choosing your next step after your degree.

Academia can feel like the natural next step after your degree. It’s familiar, intellectually stimulating, and it’s what you’ve been trained for, right?

But before you commit to a PhD, another postdoc, or start chasing the elusive tenure-track dream, here’s something worth considering:

Staying in academia just because it’s convenient is not a good reason to stay.

If you’re considering a long-term academic career, ask yourself honestly if you’re prepared for these realities:

1. It’s more about grit than genius

Academia rewards persistence, not just brilliance. Long hours, weekend work, and chasing funding often matter more than how smart you are. If you don’t want your life to revolve around work, you might feel out of sync with the culture.

2. Publishing takes priority over purpose

When it comes to securing a professorship, your success depends more on where and how often you publish than on the long-term impact of your research. If your drive comes from solving practical problems or seeing real-world results, this might wear you down over time.

3. The path is uncertain and geographically challenging

Although this depends on where you live, academic positions are generally scarce and competitive. You might need to take a junior position at a small university in a town you’ve never heard of before you can move somewhere you actually want to live.
And even if you land a professorship, only ~20% of your time might be spent on research—the rest is often spent writing grants, teaching, and handling admin work.

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This should not discourage you if you’re certain an academic career is for you—tenure tracks do still exist. But a professorship is not quite the dream we imagined when we first started studying. This realization is important, even if it’s inconvenient.

Assuming academia isn’t the right path for you, what then?

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Which Career Path Outside of Academia Is Right for You?

If you’re a science student or early-career researcher considering a career beyond academia, you’re not alone—and you’re not without options.

To help you narrow things down, we’ve organized five core career paths where science-trained minds thrive—and broken them down into concrete roles. Each section ends with a question to help you reflect on whether it’s a good fit for your interests and strengths.

Disclaimer: This is not an exhaustive list—some people land jobs as Sci-Fi Advisors. Just because a role isn’t listed here doesn’t mean it doesn’t exist. These five paths simply offer a general orientation:

1. Information and Analysis
2. Sales and Marketing
3. Research and Development (R&D)
4. Clinical and Medical Affairs
5. Business, Finance, and Policy

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1. Information and Analysis

This path is perfect if you love gathering, analyzing, and structuring knowledge.

If you’ve ever enjoyed digging through papers for a literature review, translating complex topics into digestible formats, or helping others understand new innovations—this might be your sweet spot.

There are three distinct sub-paths here:

➤ Science Writing and Editing
Turn technical information into clear and compelling content. You might write manuals, simplify academic findings, or edit research communications.
Sample jobs: Technical Writer, Scientific Editor, Communications Specialist

➤ Intellectual Property
Work on protecting scientific innovations so they can be commercialized. You’ll help register patents, evaluate the novelty of inventions, and bridge science with legal systems.
Sample jobs: Patent Examiner, Intellectual Property Analyst, Technology Transfer Officer

➤ Information and Data Management
Use your analytical skills to extract insights from data. This can range from structuring internal company databases to complex data science tasks.
Sample jobs: Data Scientist, Research Analyst, Business Intelligence Specialist

👉 Ask yourself: Do I enjoy digging into information, organizing complex ideas, and making sense of data or documents for others—without getting overwhelmed?

---

2. Sales and Marketing

If you’re energized by interacting with others, explaining technical products, and guiding decisions—this track might be for you.

In these roles, you’ll work with existing products and help communicate their value to clients, customers, or internal stakeholders. You’ll need strong communication skills and the ability to translate science into business benefits.
Sample jobs: Application Scientist, Technical Sales Specialist, Product Manager, Marketing Associate

👉 Ask yourself: Do I enjoy presenting, networking, or helping others understand the value of a product or idea?

---

3. Research and Development (R&D)

Still excited by experimentation, hypothesis-testing, and pushing the boundaries of knowledge? R&D allows you to stay close to the bench—just in a more commercial or applied setting.

Here, you’ll develop new products, technologies, or therapies that don’t yet exist. If you enjoy designing experiments and seeing tangible results, this is worth exploring.
Sample jobs: R&D Scientist, Project Manager, User Experience (UX) Researcher, Product Development Scientist

👉 Ask yourself: Do I like the idea of creating something new that solves real-world problems—even if I have to give up some of the freedom of academic experimentation?

---

4. Clinical and Medical Affairs

This is a strong option if you’re interested in translating science into medicine—especially if you enjoy cross-functional communication with healthcare professionals.

You might support clinical trials, explain product benefits to medical stakeholders, or ensure that new drugs meet regulatory requirements.
Sample jobs: Clinical Trial Manager, Medical Science Liaison (MSL), Clinical Research Associate (CRA), Regulatory Affairs Specialist

👉 Ask yourself: Am I interested in bridging science and medicine, and do I enjoy collaborating with clinicians or navigating regulations?

---

5. Business, Finance, and Policy

If you’re interested in solving large-scale problems, shaping policy, or applying your analytical mind to finance or strategy, this route offers immense impact.

It’s especially good for scientists who want to step back from the lab and work on the structural systems that support innovation, business, and research funding. Also, the pay is typically higher than in other roles—especially in the financial sector.

➤ Financial Services
Use quantitative skills to model financial trends, assess investments, or manage risks.
Sample jobs: Quantitative Analyst, Equity Research Analyst, Risk Analyst

➤ Business and Strategy
Help companies grow, restructure, or solve major organizational problems.
Sample jobs: Management Consultant, Business Development Manager, Strategy Analyst

➤ Policy and Funding
Influence science policy, funding priorities, and public research strategies.
Sample jobs: Science Policy Advisor, Grant Facilitator, Government Research Analyst

👉 Ask yourself: Am I drawn to broader, strategic thinking? Do I want to shape how science is funded, applied, or governed?

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As mentioned, other valuable career paths exist beyond the five main ones. For example, a product manager typically works at the intersection of science, business, and engineering—overseeing the development and lifecycle of a specific product, aligning it with customer needs, and ensuring its success in the market.
In contrast, a program manager focuses more on coordinating projects, timelines, and teams—often overseeing multiple related projects or research efforts to keep everything running smoothly.

If you’re drawn to broader impact, you might also consider becoming a government science advisor or working with an NGO on science policy, where you can influence decision-making and shape scientific or health policies at national or global levels.

---

Final Thoughts: What Do You Enjoy Most?

When choosing a career path, consider what kind of environment energizes you. Ask yourself:

• Do I want to work independently, or do I thrive on teamwork and communication?
• Do I want to create new things, or would I rather help others understand, progress, or improve them?
• Do I prefer fast-paced, competitive environments—or slower, detail-oriented ones?

PS: Be honest with yourself—even if your answers feel embarrassing.
For example: “I’m competitive. Success makes me happy. If I’m on a team where others drag me down, I’d rather fail alone than succeed by chance with them.”
That’s totally fine. You’re not the only one out there.

---

Most importantly, don’t feel pressured to pick just one path.
Corporate environments tend to be more flexible than academia. If you end up in a smaller company or even a startup, you’ll probably wear multiple hats anyway. 😊

Your training gave you a toolkit. Now it’s time to figure out where you’ll apply it best.

Below, you will find all resources mentioned in the talk including some additional information:

Talk Related Resources

Talk Recording
https://youtu.be/1OkfaGaSRMs

Talk Slides
http://re-advance.com/wp-content/uploads/2025/08/Integrating-Sustainability-in-Science-Safely-and-Efficiently-at-the-Leibniz-HKI.pdf

Publications

Reducing environmental impacts of marine biotoxin monitoring: A laboratory report
https://journals.plos.org/sustainabilitytransformation/article?id=10.1371/journal.pstr.0000001

A case report: insights into reducing plastic waste in a microbiology laboratory
https://www.microbiologyresearch.org/content/journal/acmi/10.1099/acmi.0.000173

Reducing plastic waste in scientific protocols by 65% — practical steps for sustainable research
https://febs.onlinelibrary.wiley.com/doi/epdf/10.1002/1873-3468.14909

Fume Hood Savings

Havard
https://sustainable.harvard.edu/wp-content/uploads/2023/09/FumeHoodWhitePaper-1.pdf

University of California / UC Davis and UC Santa Barbara
https://www.energy.gov/femp/articles/fume-hood-sash-stickers-increases-laboratory-safety-and-efficiency-minimal-cost

Additional Resources

Sustainability for Science & Data Quality

A New Approach to Making Scientific Research More Efficient – Rethinking Sustainability
https://febs.onlinelibrary.wiley.com/doi/full/10.1002/1873-3468.14736

Sustainability Guide Co-Authored by Merle Hammer

Original Version
https://tu-dresden.de/tu-dresden/nachhaltigkeit/ressourcen/dateien/campus-und-betrieb/green-lab/GoGreenGuide.pdf?lang=de

TU 2025 Verion
https://tu-dresden.de/tu-dresden/nachhaltigkeit/ressourcen/dateien/campus-und-betrieb/green-lab/Green-Lab-Guide-06_2025_ENG-1.pdf?lang=de

More helpful resources are assembled here:

https://www.linkedin.com/posts/patrick-penndorf_making-labs-greener-sharing-the-best-resources-activity-7281991446528745472-Wqkj?utm_source=share&utm_medium=member_desktop&rcm=ACoAADaG7eUBhRzKIP0RezdUPINYaEVNC7gquhc

There is a lot we can do to optimize our methods.

Often, we are only aware of a fraction of the possibilities, as technical options are usually available only to those who have worked for several years with a specific instrument.

However, there are several actions we can take that are simple in nature and will greatly benefit not only the environmental impact but also the time, money, and sample required.

I want to give you an (almost) exhaustive list of opportunities I am aware of for optimizing your HPLC workflows.

I created this list because there are plenty of opportunities, but you should be able to review it quickly to identify where you might find unused potential in your lab:

I compiled the following:

• Technical Components
• Column and Gradient Selection
• Alternative Eluents
• Approach Optimization
• Solvent Recycling

I’ll start with Technical Components & Set-up, because I think these are low-hanging fruit: the practices are straightforward, and add-ons can be quickly purchased and installed—even by non-experts.

Before we start, big shout out to Michael Meyer who significantly helped with collecting these ideas!

Technical Components & Setup for a More Sustainable HPLC

Let us start with some technical tips and gadgets that will greatly benefit your work, health and the environment:

SCAT filter units for eluent and waste bottles are an effective way to improve both safety and sustainability in the lab.

• Special GL45 screw caps with capillary feedthroughs and secure fittings prevent solvent evaporation and reduce exposure risks.
• Patented filters include a time indicator for replacement intervals, helping you change filters only when needed — avoiding unnecessary waste while maintaining safety.
• The filter’s valve design ensures no impact on system pressure.
• Filters are available in sizes tailored to all common bottle volumes.

Why it matters: Reduced solvent evaporation lowers VOC emissions and solvent waste, and using the right size filter minimizes unnecessary material use.

Protecting the Column and Reducing Waste

Inline filter frits for the eluent system protect the column from particles and contamination, extending its lifetime and reducing the need for costly replacements.

• Single-use frits are available, but reusable frits made of steel mesh or titanium can be cleaned using ultrasonic baths — a more sustainable choice that reduces solid waste.

Why it matters: Column replacements are expensive and resource-intensive to manufacture. Protecting them saves money and reduces environmental impact.

Minimizing Dead Volume with Smart Capillaries

Dead-volume-free capillary connections (e.g., Thermo Viper / nanoViper) significantly improve efficiency and sustainability.

• Enable fast, reproducible gradients, reducing run times and solvent use.
• Fingertight sealing holds at pressures above 1000 bar, removing the need for constant screwing, re-tightening, and troubleshooting leaks.
• Universal fitting design works with any column head or detector port, sealing via a PTFE ring at the capillary tip.
• Unlike PEEK capillaries, these can be used without restriction for aggressive solvents such as THF.
• Only defined lengths are available — ensuring consistent method performance and avoiding revalidation when replacing capillaries.

Why it matters: Lower dead volume and reproducible fittings improve separation quality, save solvent, and reduce the number of re-runs caused by leaks or poor connections.

Preparing the System Before Runs

Proper preparation reduces errors, prevents re-runs, and extends the lifetime of expensive components — saving time, money, energy, and solvents.

• Degas and filter mobile phases where necessary.
  Air bubbles in the solvent lines can lead to automatic system shutdowns or measurement errors. Filtration removes particles that might damage the pump or column.
• Purge all solvent channels daily, even if using the same mobile phase, to remove air via the bypass.
  Neglecting this can cause pressure fluctuations, baseline noise, or system failure during runs.
  However, purging all channels daily may be excessive if certain channels are not in use — could waste solvent unnecessarily.
• Replace flushing solvents daily to prevent microbial growth.
  Biofilm buildup can cause carryover and blockages, leading to time-consuming cleaning or costly capillary replacements. Of note, this is especially important for aqueous solvents, but purely organic phases like acetonitrile will be somewhat less vulnerable.
• Use guard columns (pre-columns) to trap contaminants from samples.
  A guard column costs ~€50, while an analytical column can range from €500–€3000 — a clear case of prevention being cheaper than cure.
• Ensure proper column equilibration with the mobile phase before starting sequences.
  Insufficient equilibration reduces reproducibility, potentially requiring re-runs (and wasted solvent, time, and energy).
• Follow calibration cycles for target compounds and confirm validity with standards (e.g., peak area calibration). This avoids both re-runs and the much worse risk of reporting incorrect results — which can waste entire project weeks.

System Care After Sequences

Post-run maintenance protects both the column and the instrument, reducing the frequency of repairs and replacements.

• After using acidic, basic, or high-salt mobile phases, flush both the column and the system with appropriate solvents before shutdown.
• Automate detector shutoff when no immediate follow-up runs are planned.
  This saves lamp hours (extending their lifespan) and reduces overall energy consumption.

Adapting Methods to Your Column Dimensions

When using a column with different dimensions from the one specified in a method (e.g., from a paper), adjustments are essential to maintain performance.

• Tools like the Restek Pro EZLC Method Translator allow you to enter both the original and your target column specifications (dimensions, particle sizes, injection volumes, gradients).
• The tool then calculates a reliable starting point for method development — reducing trial-and-error work and saving substantial time and resources.

Modern Column Selection and Method Adaptation

Why Column Choice Matters for Sustainability

Liquid chromatography consumes significant amounts of solvents, particularly methanol and acetonitrile — globally estimated at over 150,000 tons per year. Producing and disposing of these solvents has a considerable carbon footprint. Even small efficiency improvements can significantly reduce waste when multiplied across thousands of runs.

One of the most direct ways to reduce HPLC’s environmental impact is by reducing column internal diameter (i.d.) and optimizing method parameters accordingly. This cuts mobile phase use, lowers waste, and can even improve sensitivity.

Narrow-Bore Columns: Small Changes, Big Savings

• 4.6 mm i.d. → 3.0 mm i.d.
  ~60% solvent reduction per injection.
• 4.6 mm i.d. → 2.1 mm i.d.
  Up to 80% solvent reduction, ideal for LC–MS where low flow rates improve ionization efficiency.

Advantages:

• Lower flow rates = less solvent consumed and disposed of.
• Reduced stationary phase and packing material in manufacturing.
• Potential increase in sensitivity for sample-limited assays.

Considerations:

• 2.1 mm columns are more sensitive to extra-column dispersion; best used with low-dispersion systems.
• 3.0 mm columns are more forgiving and widely compatible with standard LC setups.

Translating Methods to Smaller Columns

When switching to a smaller i.d., you must scale flow rates and injection volumes to maintain performance:

• Flow rate: Scale proportionally to the square of the column radius to keep the same linear velocity.
• Injection volume: Scale to match the reduced column volume — unless a boost in sensitivity is desired.

Practical tip: Use free online method translation tools (e.g., Agilent Method Translator, Restek Pro EZLC) to calculate correct flow rates, gradient times, and injection volumes for your new column size.

Reducing column length (L) while maintaining particle size lowers analyte retention time and shortens the run. Halving the column length can cut mobile phase consumption by ~50%. If the particle size (dP) is also halved (keeping L/dP constant), the same separation efficiency can be maintained — meaning you save time and solvent without losing performance.

Benefits include:

• Shorter equilibration and re-equilibration times.
• Reduced instrument energy use (less run time).
• Faster turnaround for results and higher sample throughput.

Translating Legacy Methods to Modern Formats

Many older methods still use 250–150 mm, 4.6 mm i.d. columns packed with >5 μm particles. These can often be translated to shorter columns with smaller particles while keeping the same stationary phase chemistry to preserve selectivity and peak spacing.

Example solvent savings:

• Translating an isocratic method from a 250 × 4.6 mm, 5 μm column to a 50 × 3.0 mm, 1.7 μm UHPLC column → 85.7% less solvent per injection.
• Translating the same method to a 100 × 3.0 mm, 3 μm HPLC column (400 bar system) → 71.6% less solvent without upgrading to UHPLC.

Energy savings:

• Optimized HPLC method: 56.8% less energy.
• UHPLC method: 85.1% less energy (higher efficiency at elevated flow rates).

Very Short Format Columns

For some applications, excess separation efficiency can be traded for speed and solvent savings.

• 20–30 mm UHPLC columns can dramatically cut run times and solvent use.
• Changing stationary phase chemistry (e.g., from C18 to C18-PFP) can improve resolution enough to allow shorter columns without losing separation quality.

Example: Translating a 150 × 4.6 mm, 5 μm C18 method to a 30 × 2.1 mm, 1.7 μm C18-PFP column gave better resolution than the original method while greatly reducing solvent use.

Ultra-Short Cartridges for Targeted Analysis

With LC–MS, full separation isn’t always needed — as long as analytes are retained long enough to separate from interferences.

• 10 mm cartridge columns can reduce analysis time by 88% and solvent use by 70%.
• Efficiency drops (e.g., 6.7× lower than a 100 × 2.1 mm column), so structural isomers may not resolve — but for screening or preselection, they can be a powerful sustainability tool.

Practical Tips for Implementing Shorter Columns

• Keep the same stationary phase chemistry when translating methods to avoid selectivity shifts.
• Use method translation calculators to adjust flow rates, gradients, and injection volumes correctly.
• Consider hybrid strategies: use short columns for prescreening, reserve full methods for confirmatory analysis.

Choosing the Right Column for the Job

The right column choice can dramatically improve both efficiency and sustainability. Still:

• Standard lab setups often use 75–100 mm length columns with 2.1 mm internal diameter for general work.

• For specific applications, such as broad-range fermentation analysis, longer polymer-based columns (e.g., 300 mm × 7.8 mm) may be better — allowing full analyte separation in a single injection.
  This reduces the need for multiple runs, saving solvents and instrument time.

Tip: Always know the minimum column specifications required for your analytes. Consult:

• Experienced colleagues or application specialists from reputable suppliers (e.g., Phenomenex, Waters, Macherey-Nagel).
• Company representatives often visit research facilities monthly — arranging short on-site consultations can be very productive.

Some suppliers provide online column search tools based on analytes; in the past, mobile app solutions were discussed, though availability should be verified before relying on them.

Solid Core Particles

Solid core (superficially porous) particles improve kinetic performance compared to fully porous silica.

Key benefits:

• Higher efficiency: A solid core column with the same dimensions and particle size as a fully porous column can nearly double separation efficiency.
• Faster runs: Reduced surface area and permeability lead to lower retention times — in one example, run time dropped by 50%, cutting solvent and energy use in half.
• Improved sensitivity for impurities due to sharper peaks.
• Similar back pressure to fully porous columns — compatible with conventional HPLC systems.

Practical tip: The extra efficiency can be traded for sustainability by using shorter solid core columns (50–75 mm), further reducing run time, solvent use, and energy consumption without sacrificing resolution.

UHPLC and Particle Size Advantages

Modern UHPLC systems (>1,500 bar) support smaller sub-2 μm fully porous particles or superficially porous particles in narrow-bore columns. These allow:

• Higher efficiency separations in less time.
• Shorter gradients without loss of resolution.
• Lower mobile phase consumption.

Note: While UHPLC hardware is optimized for low dispersion, the principles of narrow-bore column scaling also apply to conventional HPLC.

Shorter and Smarter Gradients

Most HPLC users stick with the gradient they first developed — or copied from a paper — without questioning if it’s the most efficient. But gradients are one of the biggest levers for sustainability: shorten them, optimize them, and you save solvent, energy, and time every single run.

Why gradients matter

A gradient isn’t just a ramp of mobile phase composition. It determines:

• How long your run takes (and therefore how much solvent is consumed).
• How well compounds are separated (resolution vs. speed trade-off).
• How much equilibration is required before the next injection.

Unoptimized gradients often mean unnecessary “flat” baseline time, overlong separation windows, or excessive re-equilibration — all of which waste solvents and hours.

What shorter gradients can achieve

Scientific studies show that halving run times by adjusting gradient slopes can save 40–60% mobile phase per injection without losing separation quality. For example:

• Switching from a 30-minute gradient to a 15-minute gradient with optimized slope often maintains resolution but halves solvent consumption.
• Reducing post-gradient re-equilibration from 10 minutes to 3 minutes can save even more over hundreds of runs.

In practice, this means one lab running 50 injections a week could save multiple liters of acetonitrile per month — simply by trimming dead time.

2 Tips to optimize gradients

1. Remove unnecessary plateaus
   Look at your chromatogram: are there long stretches where nothing elutes? Those are prime candidates for trimming.
2. Steepen the slope strategically
   Instead of a slow, linear gradient, use segmented ramps: shallow slopes where resolution is critical, steeper slopes where analytes elute cleanly apart.

A note on regulations vs. flexibility

If you’re working under pharmacopeial methods (e.g., in pharma QC), gradient changes aren’t always allowed — regulatory frameworks often require methods to be followed exactly, even if they’re almost obsolete and wasteful. In academic or research settings, however, you usually have much more freedom. Don’t be afraid to question inherited methods: trimming gradients or re-optimizing equilibration can save solvents and time without breaking any rules.

UHPLC can make a difference Modern UHPLC systems (ultra-high pressure LC) allow the use of smaller particles and shorter columns while maintaining separation efficiency. This means gradients that once took 30–40 minutes on a 5 µm, 250 mm column can often be completed in 10–15 minutes on a 50–100 mm, sub-2 µm column — with equivalent or even better resolution, and a fraction of the solvent use.

Flow Rate Optimization

UV Detection and Flow Rate: Practical Implications

• UV detectors are concentration-sensitive: peak height depends on analyte concentration, while peak area reflects the total amount passing through the detector.
• Lower flow rates increase the residence time of analytes in the detector cell, which can increase peak area without changing the amount injected.
• Peak height changes with flow rate in more complex ways, but for small adjustments, the impact on resolution is minimal.

Sustainability takeaway:
Reducing flow rates without increasing total rune time decreases solvent use and can increase sensitivity for certain analyses, potentially allowing you to inject smaller sample volumes and shorten gradients.

Greener Alternatives to Conventional HPLC Eluents

Switching solvents is one of the most direct ways to make HPLC more sustainable – but it’s also one of the most complex. The choice of organic modifier in a mobile phase affects everything: separation efficiency, system back pressure, detector compatibility, method robustness, and even instrument wear.

For decades, acetonitrile (ACN) has been the default choice but greener alternatives do exist — from bio-based alcohols like ethanol to emerging solvents like ethyl lactate and Cyrene. While every substitution has to be well considered, if established they are much greener and in several cases even improved process and data quality. Moreover, incidents like the “Acetonitrile Crisis” showed how a bottleneck in ACN supply can limit research processes and drive prices significantly higher.

Therefore, the sections below outline realistic solvent alternatives, their pros and cons, and practical tips for implementation – so you can make informed changes without sacrificing chromatographic performance.

Why Focus on Acetonitrile (ACN) Reduction or Replacement?

Acetonitrile (ACN) is one of the most widely used organic solvents in reverse-phase HPLC due to its:

• Low viscosity → lower back pressure.
• Low UV cut-off (~190 nm) → ideal for UV detection.
• Strong elution strength in reversed-phase separations.

However, ACN has drawbacks:

• Derived mainly from petrochemical sources, with fluctuating supply and high production footprint.
• Toxic to aquatic life and requires controlled waste disposal.
• Supply shortages (as in 2008–2009) can disrupt laboratory workflows.

Reducing or replacing ACN can therefore lower environmental impact, improve laboratory resilience, and sometimes reduce costs. Below are realistic, technically tested alternatives.

Ethanol (EtOH)

Pros:

• Lower toxicity than ACN and MeOH.
• Easier disposal, biodegradable.
• Compatible with both UV and MS detection for many applications.

Cons & considerations:

• Higher viscosity → higher back pressure, especially in aqueous mixtures.
• Higher UV cut-off (~210 nm) → potential noise increase and reduced sensitivity at low wavelengths.
• May require increased column temperature to reduce viscosity and improve performance.

Practical tips:

• Use columns packed with superficially porous particles to offset high back pressure.
• Pre-heat column or mobile phase to 40–60 °C to lower viscosity.
• Works well for analytes soluble in ethanol — in some cases, analytes can be extracted directly into ethanol (e.g., UV filters from cosmetics).

Propylene Carbonate (PC)

Pros:

• Polar aprotic solvent, potential ACN replacement in RP-LC.
• Commercially available in high purity at reasonable cost.
• Produced via relatively green synthesis routes.

Cons & considerations:

• Not fully miscible with water → requires ternary mobile phases (often PC + EtOH + water).
• Higher viscosity and density → increased back pressure.
• Higher boiling point and lower vapor pressure → may reduce MS sensitivity.

Ethyl Lactate

Pros:

• Biodegradable, non-toxic, food additive status.
• Fully miscible with water and many organic solvents.
• Low cost and renewable origin.

Cons & considerations:

• Not stable under strong acidic or alkaline conditions.
• Higher UV cut-off than ACN → not ideal for UV detection at short wavelengths.
• Not widely available in HPLC-grade purity.

Example:
A mobile phase of 87% water, 10% ethyl lactate, 3% acetic acid achieved baseline separation of three pharmaceuticals on a C18 column at 60 °C in under 3 minutes.

100% Aqueous Mobile Phases

Pros:

• Non-toxic, safe, no disposal concerns.
• No UV absorbance above ~190 nm → excellent for UV detection.

Cons & considerations:

• Low elution strength at room temperature for non-polar compounds.
• Risk of phase collapse/dewetting with standard C8/C18 columns — use polar end-capped or polar-embedded stationary phases.
• Often requires elevated temperature to improve elution strength.

Cyrene

Pros:

• Produced from renewable biomass (sawdust).
• Biodegradable in <28 days.
• Low toxicity, high flash point, safe handling.

Cons & considerations:

• Immiscible with water — requires ethanol or another co-solvent.
• High viscosity and boiling point (227 °C) → high back pressure, possible MS limitations.
• UV detection below 350 nm problematic due to background noise.
• Not currently available in HPLC-grade.

Example:
In RP-HPLC, ternary mixtures of Cyrene + ethanol + buffer allowed separation of metronidazole and moxifloxacin at 50 °C on a monolithic column with acceptable back pressure (~130 bar).

Practical Strategies for Using Alternative Eluents

1. Start with partial replacement (e.g., replacing 20–30% ACN with EtOH) to evaluate performance.
2. Adjust temperature to manage viscosity-related back pressure.
3. Use compatible columns — superficially porous or monolithic formats often handle viscous solvents better.
4. Validate sensitivity impact — higher UV cut-off solvents may require detection at higher wavelengths or MS-based detection.
5. Test method robustness — alternatives may affect retention time stability under small variations in pH, temperature, or composition.

Smarter Method Development

• Dry-run gradient simulation: If your software allows, simulate gradients before running them to spot obvious inefficiencies.
• Start isocratic where possible:
• Tip: Start Isocratic Where Possible
  • If your separation doesn’t need a gradient, isocratic runs are almost always more sustainable.
  • Lower solvent use: Composition stays constant, so you avoid the long high-organic washes and re-equilibration phases that gradient methods require.
  • Shorter equilibration: Switching from one run to the next is faster because the column is already at the correct mobile phase composition.
  • Easier solvent recycling: Isocratic methods make post-detector recycling straightforward.
  • Practical check: If your analytes elute within a narrow retention window in a gradient, try an isocratic method with that composition — you may get equivalent resolution in less time, using less solvent.
• Short equilibration: Validate the shortest possible re-equilibration time — many default methods leave unnecessary idle times between runs.

Optimizing Sample Preparation

When people think about making HPLC greener, they usually start with the instrument — smaller columns, alternative solvents, clever fittings. But sample preparation is often the hidden giant in both time consumption and environmental footprint. It’s easy to overlook because it feels like “the routine bit before the actual analysis.” Yet, if you optimise your prep, you can save hours of work each week and cut waste dramatically.

Why optimising it matters:

• Sustainability: Less solvent, fewer consumables, and smaller hazardous waste volumes.
• Time savings: Trimming just 2 minutes per sample in a 100-sample sequence means over 3 hours gained — enough to run an extra sequence or wrap up earlier.
• Data quality: Cleaner, more consistent samples mean fewer failed runs and less rework.

Practical changes that deliver:

• Miniaturise volumes: Use micro-extraction techniques (e.g., SPME), smaller vials, and scaled-down filters.
• Direct injection: If your matrix is clean enough, skip concentration/dilution steps.
• Batch preparation: Prep all daily samples at once to avoid repeated purging, solvent refills, and cleaning cycles.
• Greener solvents: Swap chlorinated or high-toxicity solvents for ethanol or ethyl acetate when possible.
• Smarter filtration: Pre-centrifuge to avoid filter clogging; filter only when necessary.

Using Combination Method

If you plan to use an HPLC–MS (LC–MS) approach, there are often opportunities for savings that go beyond just getting great data. LC–MS setups are already among the most sensitive analytical systems available — often 10–1000× more sensitive than UV detection — which means you don’t always need the long columns, high flow rates, or large sample volumes that standard HPLC methods rely on.

Why This Matters for Sustainability

• Smaller sample amounts: Lower detection limits mean you can inject far less sample — good for precious, hard-to-make, or hazardous materials.
• Shorter columns and runs: Since MS can identify compounds by their mass, you don’t always need full baseline separation, so you can shorten gradients or use smaller formats.
• Less solvent waste: Lower flow rates directly reduce mobile phase use and disposal volumes.

Capillary HPLC for LC–MS

Capillary HPLC reduces column internal diameter to 100–500 μm, running at 0.4–100 μL/min instead of the 1–2 mL/min of a typical analytical column.

In practice, this means:

• Over 90% less solvent per run.
• Sharper peaks for the same mass injected — giving better sensitivity without extra sample prep.
• The ability to run dozens of injections before you’ve even used as much mobile phase as a single standard HPLC run.

Solvent Recycling: Turning Waste Back into Usable Mobile Phase

In isocratic HPLC, the mobile phase composition stays constant throughout the run. This means that the solvent exiting the detector is identical in composition to the one entering the column — apart from minor changes due to analyte and impurity carryover.

This makes it straightforward to collect the “clean” post-detector solvent and feed it back into the mobile phase reservoir after appropriate filtration and (if necessary) degassing.

In gradient elution, however, the mobile phase composition changes over time, so the effluent is constantly varying in proportion of organic and aqueous components. Recycling would require real-time composition matching — impractical in most lab setups and potentially risky for method reproducibility.

Solvent Recycling Systems

Basic principle of solvent recycling can summarized as: divert the post-detector solvent to a collection system.

There are two main approaches:

A. Manual Recycling (No Computer Support)

• Effluent is collected in a clean container during known “blank” parts of the chromatogram — typically before the analyte of interest elutes.
• This requires knowledge of retention times and precise timing of collection to avoid contamination.
• Solvent is then filtered (0.2 μm or finer) and reintroduced to the mobile phase reservoir.

Pros:

• Low cost, no specialized equipment.
• Easy to implement for single-analyte methods with stable retention times.

Cons:

• Requires careful manual operation.
• Not suitable for complex mixtures with unpredictable peaks.

Automated Recycling (Computer-Controlled)

• Integrated with the HPLC’s data system or an external controller.
• The detector signal triggers a divert valve that sends the flow either to waste (when peaks are eluting) or to the recycling reservoir (during baseline).
• Many systems allow the user to set a threshold absorbance value — when the signal drops below this, solvent is automatically routed for recycling.

Pros:

• Reduces human error and contamination risk.
• Ideal for high-throughput labs running the same method repeatedly.
• Can recover a significant percentage (often >80%) of mobile phase for reuse.

Cons:

• Requires investment in hardware and software integration.
• Still best suited for simple isocratic methods with stable baselines.

Practical Considerations for Safe Recycling

• Filter carefully: Use high-quality solvent-resistant filters (PTFE or PVDF) to avoid introducing particulates back into the system.
• Monitor solvent quality: Over multiple cycles, trace impurities can accumulate — periodically replace all solvent to maintain method performance.
• Use compatible containers: Ensure collection and storage vessels are chemically resistant to your mobile phase.
• Degas before reuse: Prevent bubble formation in pumps and detectors.

Cell culture and S2 work is always a hot topic.

Since it involves culturing cells and infecting them, the question naturally arises: what can and cannot be done sustainably in these environments?

The answer is – a lot. Even in S2 labs, significant amounts of plastic waste can be saved. I described one of my own workflows in detail in this publication (“Reducing plastic waste in scientific protocols by65% — practical steps for sustainable research”), but here are some key principles for routine work and those I have worked out with Jakob.

First, many doubt whether waste separation is even allowed. Legally, it is. The German regulation for genetically modified organisms explicitly states that waste can be discarded normally — and therefore recycled — if it has not been contaminated. The simple rule is this: everything that didn’t come into contact with genetically modified or hazardous materials can be separated.

That means, for example, the plastic and paper wrapping of serological pipettes can go into recycling rather than the biohazard bag.

Beyond disposal, there are small but impactful tweaks in daily handling. If you are splitting one cell line into several flasks, you can reuse the same serological pipette to distribute medium across them. Choosing larger pipettes when working with only one cell type – for example, dispensing PBS or trypsin in one go – also reduces pipette use. For some cell lines, culture flasks can even be reused for routine maintenance, provided you check carefully that the cells remain healthy. And when working with small flasks, adding medium first and then cells means one serological pipette can be used for both steps.

Safety, however, is non-negotiable. Jakob, for example, never reuses serological pipettes or shares them between parallel splits when handling two different cell lines. This eliminates any risk of cross-contamination, which would be nearly impossible to detect later if a few stray cells started growing in the wrong culture.

As always, you should only go as far as you feel comfortable. Implement controls, pay attention to best practices such as expelling liquids completely, and above all, gain sufficient experience before attempting to adapt. Experience is what allows you to judge whether a practice is safe or not. Even then, the transition should be gradual. Make small changes step by step, so that you can keep your flow without becoming insecure, and so you can trace what works well and what doesn’t.

And remember: if you are in a hurry, don’t force sustainability into the workflow — that’s when mistakes happen. Finally, culture matters. How do you communicate what you expect from your lab members? Make sure postdocs and PhDs are aligned and that sustainable practices are included in training. When new colleagues join, take the time to let them shadow for two or three sessions so they learn the lab’s culture of working. Having someone look over a shoulder is often enough to correct habits, catch overlooked opportunities for optimization, and make sustainability part of the lab’s shared standard.

Jakob investigates how different Candida albicans strains cross an intestinal epithelial barrier grown in transwell inserts.

The aim is to test whether the presence of a wild-type, strongly hyphae-forming strain increases the ability of a clinical isolate to translocate and cause damage.

It’s a resource-intensive workflow, involving infection steps, barrier measurements, and potential downstream analyses.

And this was the perfect example to show how experimental design can optimize both data acquisition and environmental footprints.

In other words, this shows that if we optimize our workflows, we often become more sustainable at the same time.

Let’s go through how this looks in practice:

The workflow begins with preparing transwell plates seeded with a 70:30 mix of C2BBe1 and HT29 cells. All you need to know: two intestinal cell lines.

Of note, these transwell inserts allow one to check for barrier integrity since any damage allows particles or in that case fungi/spores to pass through.

Thus, once the epithelial barrier is formed, Candida strains are added alone or in combination. The core of the experiment is to measure barrier integrity.

There are four main ways to measure barrier integrity:

• FITC-Dextran assay: A permeability test where fluorescently labeled dextran (FITC) is added to the top of the epithelial layer. If the barrier is leaky, FITC-Dextran passes through to the bottom chamber. Measuring fluorescence with a plate reader shows how much leakage occurred.

• TEER (Trans-Epithelial Electrical Resistance): A direct, non-invasive method to measure barrier integrity by placing electrodes on both sides of the transwell and recording electrical resistance. A tight, intact barrier gives high resistance; a leaky one gives low. It takes only minutes and doesn’t disturb the cells, so samples can still be used afterward.

• LDH assay: Measures lactate dehydrogenase, an enzyme released into the supernatant when cells are damaged. By comparing LDH levels to a standard curve, researchers can quantify cell death or barrier disruption. It’s a biochemical readout that complements TEER’s functional measurement.

• CFU plating (colony counting): To measure how many Candida cells actually cross the epithelial barrier, samples from the bottom compartment are collected, treated with Zymolyase to break fungal cell walls, and then plated on agar in serial dilutions. After incubation, each colony that grows represents one viable fungal cell (colony-forming unit, CFU). Counting these colonies gives a direct measure of translocation and fungal survival.

Here is what Jakob decided to do:

Instead of relying on FITC-Dextran assays — which are commonly used in other labs as the easy go-to but require specific reagents, extra incubation time, and a plate reader — he chose TEER measurements. TEER can be combined with LDH and CFU plating, is direct, takes only minutes, and doesn’t disturb the cells.

By using TEER first, he avoids an additional incubation step, cuts reagent costs (and their environmental impact), and eliminates the plasticware that would have been needed for FITC-Dextran. And because TEER leaves the cells intact, he collects supernatants for LDH assays or other readouts.

Why not run FITC and LDH together? For LDH assays, they collect 50 µL of supernatant from each transwell insert – half of what’s available – to avoid disturbing the barrier. This means LDH and FITC-Dextran can’t be run from the same well in the same experiment.

Colony-forming unit (CFU) plating requires Zymolyase digestion — a two-hour step. The team avoids combining this with FITC-Dextran, which would extend the experiment by another hour and require extra consumables. Instead, they keep workflows separate unless both readouts are absolutely essential.

For us, this seems obvious, but it requires taking a step back and deciding which assays to prioritize. Once again, optimizing data acquisition and quality leads to the same conclusion as sustainability considerations. The critical step is to sit down and search for alternative methods at the level of experimental design.

But experimental design stretches beyond methodology. Jakob also looked closely at the design of single-use items and pipetting orders.

For example, instead of using 1.5 mL conical-bottom tubes for washes, they work with flat-bottom 2 mL tubes. The flat base allows pouring off washing medium without disturbing the pellet, meaning no pipette tip is needed. The same shape also makes vortexing more effective for resuspension.
What looks like a small change eliminates several plastic tips per wash step and saves time at the bench.

In LDH assays, they reuse a pipette tip for both technical replicates when handling low-risk reagents like stop solution, cutting tip use in half for that step. Reservoirs for non-hazardous solutions are labeled, rinsed, and reused rather than discarded after one use.

At the same time, he is careful to use fresh tips for dilution series, pipetting from higher to lower concentration to avoid cross-contamination — again showing that sustainability never overrides data integrity.

Moving to CFU plating, technical replicates are plated in duplicate for the same reason. This doubles the number of plates, but it catches variability early and prevents entire experiments from being repeated. Supernatants are also kept in the fridge for short periods in case re-plating is needed — another safeguard against wasting days of work and materials.

Sometimes experimental design cannot optimize sustainability. In his plate layouts, Jakob avoids the outer wells because, due to temperature differences and humidity variation, medium evaporates faster and cells grow more slowly.
That also means differentiation takes longer. This would disrupt workflows. If possible, the number of replicates is adapted; otherwise, empty wells are accepted as an unfortunate reality.

Finally, a few smart tweaks stand out. For cleaning the electrode during TEER measurements, they keep two 50 mL tubes — one for PBS, one for ethanol — and reuse them for up to a month. Ethanol volume is kept slightly lower so it never contaminates the PBS.

I spent two days with Dr. Jana and Dr. Evelyn Molloy in in their lab, where most project evolve around Molecular Biology workflows to prepare samples to extract biomolecules for HPLC analysis.

We walked the lab from one end to the other, following the natural flow of experiments – from preparing media to handling waste – looking for where they already save resources and where small tweaks could go further.

What I noticed quite quickly was sustainability here wasn’t a “special project.” Many of these practices are embedded in the daily workflow. The trick is recognizing them, optimizing them, and making them part of the lab’s shared culture.

Instead of providing a linear walkthrough, I decided to highlight two main aspects that I think will help you most in understanding sustainable practices: teamwork and creativity.

How they Create A Culture of Sustainability through Teamwork

What stood out most during my walkthrough wasn’t just the practices themselves, but how they reinforce each other to create a lab culture where sustainability is the norm.

The team has found a balance: some things are decided collectively so everyone benefits from a common default, while other choices are left to individuals.

Nevertheless, with Jana and Evelyn as a driving force, they were able to bring about large-scale change that involves several people. Moreover, once sustainable practice is culturally anchored, new lab members and students will automatically adopt these best practices. It’s like a snowball that grows and gathers momentum rolling down the hill – and it was set in motion by just these two individuals.

Alright, let me start where the basics of many of their experiments begin – media preparation.

Agar and yeast extract are bought in bulk and portioned into smaller reusable containers when needed. Simple but effective. Once storage space has been defined, it saves money and generally reduces the risk of not having sufficient supplies.

And once a lab is set up for bulk, it naturally extends to other areas like supplements and buffers. Especially if a staff member or single postdoc is handling procurement and restocking, such a culture shift can make a big difference.

Remaining in the media prep area for a moment, they showed me that Falcon tubes used to prepare vitamins or minerals are washed and reused – before adding them to the media, the contents are sterile-filtered, so reuse doesn’t add risk.

Where possible, glass replaces plastic entirely. The fact that washed tubes and glass bottles are available nearby makes the sustainable option the convenient one, and convenience drives uptake.

Of course, switching to glassware is often only possible if sufficient team members are on board, since autoclaving and cleaning have to be coordinated – but this is what they pulled off. And once implemented, it is one of the best ways to reduce waste while building a new common standard for the entire lab.

Choices made at the system level often help anchor sustainable practices. Jana walked me to the thermocyclers: they are set to idle at 12-14 °C instead of 4 °C. It’s a safe, energy-saving default.

Nobody is prevented from changing it, but very few do. Think about organ donation: if you have to opt in, very few do; if you have to opt out, very few do too. Once the baseline is sustainable, cultural change is much quicker. That means small structural changes can ripple across the whole lab without requiring active effort from each individual.

Then, a really big surprise for me followed, but one that perfectly demonstrates how teamwork saves waste. In their lab, agarose gels are reused until they’re fully consumed. If lanes remain unused, gels are kept in buffer for up to five days so someone else can finish them.

In quick control experiments, gels are even “combed from both sides,” doubling their utility. They know that many have similar conditions to run, so gels (often at 1%) are kept.

One person’s frugality becomes a resource for the next, which saves a whole lot of time for each individual. Gel buffers are swapped roughly once a month, and when evaporation occurs, only water is topped up. These are judgment calls made by individuals, but they fit into the broader culture of using what’s needed and no more.

Energy-saving practices are handled with the same pragmatism. Incubators at 35 °C and 37 °C stay on to ensure constant availability, but shaking functions are turned off when not in use.
This makes sense because the team agrees on what absolutely needs to be running, while individuals are responsible for switching off functions they don’t require. During weekends or holidays, switching those off is often feasible too.

Later, I was accompanied by Evelyn. I noticed that she didn’t wear gloves. She told me that no one really did. They knew which processes were being run, and those didn’t need gloves in that area. Where ethidium bromide is used, areas are clearly indicated, and when hazardous substances are handled, gloves are available nearby.

Here, we also find an important benefit of open communication and optimization: where EtBr is handled, students are taught right away- one glove for the handling hand, the other left bare to avoid cross-contamination to tubes or items. The point is that only if you trust your colleagues will you work efficiently as well.

Finally, as mentioned, they took care to leave everyone a personal choice. Again, psychology influences these habits. Glass serological pipettes are stored right next to the plastic ones after autoclaving. Nobody is forced to use glass, but by making the reusable option equally visible and accessible, more people reach for it. And if plastic is ultimately necessary, everybody knows where to find it.

All in all, to my mind, they have shown that establishing a culture around sustainability is extremely powerful – it enables practices that individuals cannot achieve alone, such as a complete switch to glassware, but also nudges better practices like changing PCR holding temperatures.

Nevertheless, to my mind, it is just as important that through establishing such a culture, most will save significant amounts of time, and safety will become a topic openly discussed.

Creativity as a Driver of Sustainable Lab Practice

When we think of creativity in the lab, we normally relate it to cool experiments or interpreting confusing results.

But creativity also plays a crucial role in shaping how we work.

Many of the most effective sustainable practices don’t come from new products or expensive equipment, but from rethinking the basics: asking whether the “standard way” is the only way, and trying something different.

Jana and Evelyn have adopted several really creative approaches. How they go about it? First, by being open-minded and reconsidering whether they could reduce. Second, by asking whether it was safe and, if necessary, coming up with a small test to control for changes.

When I was with Jana, she and her student were about to plate bacteria. Every microbiologist knows the odd frustration of throwing away a sterile plastic spreader after just one use…

Here, they bend the end of a pipette tip instead. It’s sterile, fast, and it avoids an entire category of single-use items.

The same mindset shapes how they prepare competent cells and pick colonies. Glycerol stocks are diluted only to the amount needed -1:10 in most cases, 1:2 – 3 when needed. Toothpicks are used from both ends before disposal.

Tiny changes, yes, but the habit of questioning “Do I really need more?” reinforces itself across hundreds of small actions.

I worked with Jana through an entire PCR workflow. These setups also benefit from this openness to new approaches. If multiple reactions use the same template, DNA is diluted in one well and split across the reactions – reducing pipetting and ensuring equal concentrations.

Mixing samples with loading dye is done on washable mixing plates or parafilm, rather than fresh tubes. Even PCR plates are reused (with silicon lids from Starlab) after thorough cleaning. Many labs would dismiss this outright, but here they tested it, validated it, and adopted it where it works (not for qPCR, but perfectly fine for qualitative checks).

At the gel bench, they challenge one of the strongest “lab dogmas”: one tip per sample. Instead, the same tip is used for 30–60 samples when they only check bands.

With careful rinsing or complete expelling, it works without contamination problems. Again, it’s not carelessness – it’s rethinking whether the old rule was ever necessary for this application. If you have 60 samples, it also saves a lot of time – making both you and your PI happy.

Ligation and assembly kits are another place where imagination pays off. Instead of following the recommended 20 µL reaction volume, they routinely work with 10 µL, sometimes even 5 µL for Gibson or NEBuilder assemblies. Since downstream transformations only use 2 µL anyway, this avoids unnecessary reagent waste. And if it doesn’t work the first time, the smaller reaction makes it easier to simply repeat.

Do I need to mention that your group leader is going to love you for the money you save them?

One reason I was so excited to work with these two was that Evelyn mentioned they reused their electroporation cuvettes. I was intrigued. Admittedly, I only used electroporation during my university years, but I had never heard of someone reusing these cuvettes. Because of their contact with bacteria and the metal included, they are often autoclaved and landfilled, never recycled.

But these two questioned the common narrative. They reuse them more than 30 times with a defined cleaning routine: rinse with water, wash with ethanol, rinse again, rinse twice with deionised water, air-dry, and UV-sterilise for 20 minutes.

Again, this was thought up, tested, and eventually established.

OD cuvettes get different treatment – washed only with water, since ethanol degrades them. But they are reused as well.

Take the filtration station. Instead of just accepting the default, they compared setups. A Steritop filter top weighs 125 g, with a plastic bottle including lid worth 89 g. Both are single-use.
But noticing the waste, they found a filter top in which the filter unit could be switched (purchasing the filters from Merck) while using a normal glass bottle to collect the prepared solution/medium. That means all that is thrown away now is the little filter.

Over a year, that small difference adds up to kilos of plastic saved. It’s not a new invention—it’s looking at the same equipment with a different lens.

I want to highlight once more that safety was always on their minds.

Let’s return to what we talked about already: glove use. They don’t wear gloves for every single step. For ethidium bromide gels, hazardous chemicals, or S2 organisms – yes, gloves are non-negotiable. But for routine E. coli hood work or pouring medium, no gloves. This isn’t about ignoring safety – it’s about applying real risk assessment. The benefits go beyond waste reduction: less skin irritation, better dexterity, and a clearer focus on careful handling.

What ties all of this together is not a single piece of equipment or policy – it’s the willingness to question defaults

Creativity here means asking: Is this rule still valid? Can I do it differently without compromising safety or results? Sometimes the answer is no, and the standard practice remains. But often, the answer is yes—and the result is a lab that saves time, money, and waste while teaching its members to think critically about every step.

Reducing waste

Reducing plastic waste within the laboratory can be achieved by:

• Optimizing protocols to prepare solutions in a single instead of separate tubes
• Using items in a smart manner by rethinking their application possibilities (e.g., mixing loading dye and samples for agarose gels after kit use on a piece of parafilm or within a reusable mixing plate instead of separate tubes)
• Using alternatives such as glass or metal items for flasks, dishes, and serological pipettes)
• Minimizing the size of consumables (especially tubes, serological pipettes, pipette tips to the required volume, potentially pipetting twice)
• Optimize for the least necessary conduct (e.g., leaving out parafilm when evaporation of fluid is not an issue)
• Pouring solutions where precises volumes are not decisive (e.g., washing steps)
• Reusing Falcon tubes, potentially after rinsing, especially for frequently used solutions (e.g., Tris solutions when preparing SDS-PAGE gels)
• Reusing pipette tips, tubes where cross-contamination is not an issue (e.g., pipetting samples in agarose gels to control for restriction digests)
• Reusing items (e.g., electroporation cuvettes or tissue/cell strainers after cleaning, cuvettes for OD measurements, weighting boards)
• Precise calculation and bulk preparation of reagents and solutions
• Automatization and miniaturization through automated systems which pipette more precisely and are less error prone
• Conscious use of gloves (see our previous lesson to learn all you need to know)
• Using items to their capacity (e.g., maximize use of wells in a 96‑well plate, all targets on an MS plate or both ends of a toothpick to pick colonies instead of pipette tips)
• Sharing items and resources to their maximum potential (e.g., parts of unused agarose gels can be kept in buffer for up to 1 one week to be used for general PCR or restriction digest controls)
• Purchasing only what is necessary (tubes without caps, DNA/RNA extraction kits without collection tubes)
• Making use of take-back programs for plastic items, including Styrofoam
• Separating waste into plastic, paper, others, autoclavation (e.g., plastic wrapping from serological pipettes tips can be discarded in plastic and paper after separating instead of autoclavation waste bins)
• Consider reuse instead of discarding (Polystyrene boxes can be used as ice boxes or to fill empty spaces in cold storage, Packaging can be used to send reagents or samples to other laboratories, Polystyrene can be used to store glass items safely)

-> Cool Examples from my consulting:

The idea of using a pipette tip to spread colonies was passed down from an Asian scientist to a postdoc, just as it was to me. Pro tip: it works best with a P200 tip – a P1000 is often too large.

The same group reused their electroporation cuvettes up to 50 times – basically until the plastic turned yellow and brittle. Their washing procedure: 1× water (to prevent DNA precipitation) → 1× ethanol → 1× water → 2× desalted water → air drying → UV for 20 minutes.

Improving experimental conduct and design

• Proper experimental planning can be achieved by:
  • Leveraging existing literature to avoid redundant experiments
  • Robust statistical planning (especially power analysis to assure interpretable outcomes) help reduce sample sizes and enhance statistical validity (e.g., group sequential design for work entailing animals)
  • Carefully chosen experimental conditions with proper controls (e.g., using painkillers with known mechanism of actions for control conditions)
  • Reducing use of animals by switching to in vitro / in silico or smart experimental design (e.g., reducing breeding through smart knock out/in strategies)
  • Re-optimizing protocols that were tailored for a specific tissue/sample/chemical
  • Reviewing possibilities for optimization in consumable utilization ahead of conduct (including material, size and number of consumables)
  • Exploring preparation procedures (e.g., optimizing pipetting schemes and master-mixes to reuse tips and tubes)

• Optimization of experimental produces is possible through:
  • Adopting safer and more benign alternatives for commonly used reagents in experiments (e.g., DNA staining solutions, microscopic slide mounting agents, lysing agents, or protease inhibitors)
  • Exploring alternative experimental approaches (e.g., using Supercritical fluid chromatography to avoid organic solvents needed in HPLC or modes of inducing cell death)
  • Applying combination methods such as LC-MS to minimize sample preparation and maximize data acquisition
  • Switching to miniaturization procedures (automated pipetting, solid phase microextraction (SPME))

• Best practices for experimental conduct include:
  • Considering potential for downstream use or regeneration (e.g., regeneration of nucleic acid extraction columns)
  • Minimization of experimentation (e.g., reducing PCR volumes to 10uL instead of 20uL if the overall presence of a construct or the success of a digestion should be assessed)
  • Implementing strategies and frameworks to ensure best practices (e.g., handling pipettes upright when pipetting)
  • Awareness of toxicity of reagents in use for proper handling and discarding (e.g., including closing lids to avoid evaporation)

• Impact can be reduced by initiating collaboration with:
  • Colleagues in co-preparation of solutions, sharing of samples or co-use of machines (e.g., water baths)
  • Other groups to share equipment
  • Core facilities or partners to avoid unnecessary establishment of new methods

-> Cool Example from my consulting:

One of the scientists decided to use larger 2 mL tubes instead of 1.5 mL tubes. The reason: since the 2 mL ones have a round bottom instead of the conical shape of the 1.5 mL tubes, he could resuspend the pellet by vortexing without needing a pipette. Additionally, he could pour off the supernatant after centrifugation without needing to pipette it out, since the larger surface ensured the pellet adhered to the bottom. In doing so, he saved two tips and quite a bit of time.

Changing procurement and purchasing processes

• Planning orders carefully by:
  • Creating an internal system to track chemical inventory and consumable supplies to minimize unnecessary orders
  • Reduce frequency of orders
  • Collaborating with other laboratories or facilities to collect orders
  • Contact manufacturers to organize more efficient delivery and packaging

• Choosing products consciously by:
  • Procuring items in quantities aligned with future usage (i.e., not more than necessary but in bulk if possible)
  • Purchase large quantities (e.g., NaCl or bacterial/fungal medium components)
  • Emphasizing sustainable packaging practices, favoring minimal material usage and biodegradable materials where possible
  • Opting for specific shipping methods and alternatives to conventional cooling methods (e.g., when ordering polymerases without cooling or oligos dry)
  • Choose products of lighter weight (e.g., gloves, tubes, pipette racks)
  • Choose products that have exchangeable parts (e.g., vacuum filters in which just the filter is exchangeable instead of the entire plastic housing is single use)
  • Thoroughly evaluating feasible alternatives based on certifications, life cycle analyses, and sustainability practices

• Choosing the optimal supplier:
  • Preferring local suppliers to reduce transportation-related emissions and dependency on global supply routes
  • Preferring certified suppliers and articles
  • Exploring take-back programs and consider second-hand purchases to enhance sustainable procurement practices

-> Cool Examples from my consulting:

To prepare their medium, a group originally used a plastic filter top (125 g) and a dedicated plastic bottle (75 g), both of which had to be discarded after a single use. Recently, they found a reusable filter top where only the filter itself needs to be replaced — they simply buy the correct diameter and pore size from Merck and use glass bottles for filtration.

They also showed me a very useful 96-well PCR plate with a silicone sealing mat, allowing both pieces to be washed and reused for routine PCR checks.

Reducing paper, water and energy use

Paper

• Transition to digital sources like electronic lab journaling and online publications
• When printing is necessary, using recycled paper and opt for double-sided printing on previously used paper
• Practice conscious use of wipes (i.e., avoid stacking of unnecessarily many wipes)
•  Reuse wipes (e.g., when drying slides)

Water

• Minimize water use, for example by soaking steps and mechanical cleaning
• Consciously discern water types (tap, distilled, double distilled etc.)
• Use only as much ice as needed
• Ensure proper maintenance of equipment (e.g., filters of reverse osmosis systems)
• Choose proper equipment and use (e.g., medical grade sterilizers are often designed to run 24/7, exceeding needs in common laboratories and Switching from traditional steam-jacketed autoclaves to newer types without steam jacket)
• Install Low-Flow Aerators
• Reusing “Wastewater” (e.g., institution wide when high grade water is reused for less curcial tasks or with counter-current rinsing principles)
• Choose waterless equipment (e.g., waterless condensors)
• Avoid Single-Pass Cooling Systems by using Closed Loop Systems
• Consciously choose batch-type washers versus tunnel washers depending on your usage
• Consider automated systems to provide only the necessary amount of water for animals, reducing waste while maintaining animal welfare.
• Install high-efficiency wet scrubbers that minimize water use while effectively controlling emissions
• Reduce the frequency and volume of hood wash downs by adopting targeted cleaning practices and evaluating actual cleaning needs
• Implement water recycling and treatment systems to reduce the overall water demand of cooling towers
• Optimize settings and capture HVAC condensation water for non-potable purposes, such as irrigation or cooling tower makeup water
• Actively search for and report leaks in taps and pipes

Energy

• Regularly organizing and cleaning digital inboxes to prevent unnecessary data storage
• Maintain a tidy system for experimental data, avoiding unnecessary duplication and keeping a safety copy securely stored on a hard drive
• Exercise caution with AI technologies and use of search engines due to their potential high energy consumption
• Evaluating the necessity of video in online meetings and switch to audio-only when possible to minimize data and energy usage
• Keeping laboratory fume hood sashes shut and turn machines off when not in use (e.g., water baths)
• Double check A/C settings and location of thermostats to avoid unnecessary cooling due to exhausted air
• Reduce air exchange frequences in HVAC systems if possible
• Set up an organization software for people to coordinate use & turn on/off of equipment
• Search for new innovations and energy efficiency when purchasing new equipment (i.e., lower energy usage e.g., in freezers or HPLC systems, choosing LED instead of halogen using microscopes)
• Create concrete plans/responsibilities for light & equipment turn off during weekends and vacation
• Optimize equipment placement (e.g., move freezers away from walls to allow for proper heat dissipation and consider how many and which pieces of equipment shall be placed in a single room)
• Setting PCR-Holding Temperature to 12°C or higher

Optimizing Equipment Use

• Choose instruments with reference to:
  • Lifetime (e.g., photomultiplier tubes have longer lifetimes)
  • Capacity (e.g., volume of sterilizers & autoclaves)
  • Components (using low-boiling-point solvents in air-cooled condensers to reduce energy consumption)
  • Minimizes reagent use (e.g., Nitrogen consumption in MS or HPLC columns with smaller inner diameter to reduce solvent consumption and waste creation)
  • Performance (necessary precision, combination methods such as LC-MS reduce sample preparation)
  • Running mode (e.g., enables change to more sustainable alternatives (Hydrogen as carrier gas instead of He in GC/MS)  or enables internal reuse (e.g., automated recycling of the mobile phase for example after absorption of the impurities)

• Making a conscious choice about what methodology to use (e.g., wet vs dry blotting, on site analysis, high throughput analysis, combination techniques such as LC-MS)
• Exercising best practices (e.g., not let elution fractions from chromatography columns evaporate or using all spots on matrix array plates for MS, putting as many samples on one microscopy slide as possible)
• Being aware of the robustness of methods (e.g., ability to reuse TLC capillaries after rinsing)

• Reducing energy use by:
  • Developing an energy plan, i.e., when to turn on and off individual machines
  • Using strategies like multi-plugs to turn off ovens and water baths during inactivity or employing smart plugs for automated on/off cycles
  • Considering carefully how you use equipment (settings including scanning area in microscopy)
  • Modifying freezer temperatures, such as increasing from -80 to -70
  • Using covers for water baths and replace oil baths with more efficient alternatives like metal heating blocks or efficient oil pumps
  • Operating dishwashers and autoclaves only at full capacity
  • Consciously choosing levels for the A/C set-up
  • Choosing sufficient settings (e.g., temperatures and shaking frequency in incubators
  • Disconnect ducts of fume hoods in which non-toxic chemicals are use
  • Hibernate no longer used fume hoods in order to save energy since they are normally included in the general ventilation system
  • Hand over, use secondary market or donate equipment that is no longer in use

• Reducing water use by:
  • Implementing low-flow aerators to conserve water
  • Using closed-cycle cooling systems and waterless liquid-cooled condensers with low-boiling-point solvents as an alternative to single-pass cooling methods

Optimizing waste treatment

• Making sure that evaporating waste is handled properly (e.g., stored in a hood or closed container)
• Using old jerry cans/flasks/container as waste containers (or already contaminated tubes)
• Create a plan how to handle cooling packs, animal bedding, Styrofoam etc
• Establishing education and indication systems (e.g., easily undersandable stickers on waste bins, using and a database with necessary educational resources)
• Repairing broken glassware (e.g., through chemistry faculty internal gaffers) and old pipettes
• Proper separation of waste (e.g., paper vs plastic, contaminated vs not-contaminated)
• Identify biosafety and contamination sources to avoid unnecessary contimation (e.g., in some institutions, all items touching a bench count as contaminated, thus, de-wrapping packaging outside the lab avoids autoclavation before recycling)
• Separate items early on (e.g., when caps of tubes are not used, they can be discarded in common plastic trashes)
• Create robust recycling schemes in cooperation with 3rd parties (i.e., developing separation schemes in HDPE vs PP that actually are maintained in transport and recycling by the waste handling parties)
• Ensure buy in from all stakeholders when it comes to waste separation (especially cleaning staff and waste handling parties must be informed and screened)
• Try to compost biological materials or animal bedding