Hello Reader, how do we identify sustainable practices?
As you might know, I work as an advisor for universities, institutes, and companies.
Lately, I came across a prime examples of how we can drive change using three simple tactics.
Let’s discover what these are and how you can copy-paste them:
Today's Lesson: Tactics To Find And Implement
How we can identify and safely drive change in labs
Number Of The Day
More than 50 years ago - in 1971 to be precise - a method was proposed that can increase viability, detection, and time efficiency when growing bacteria. Moreover, as it would turn out, this technique can save more than 80% of plastic waste compared to standard methodology. However, fast forward to today, only few laboratories employ this technique. Therefore, let’s find out why and how to change that safely.
1971
Towards More Sustainable Practice
One of the most common missed opportunities for sustainability resides in the literature.
While Sharpe and Kilsby described the idea of drop plating as early as 1971, you can find papers such as the one in the centerthat offer further ideas for optimization (published in 2003). The paper on the left, from 2020, demonstrates how using this technique can lead to significant time, resource, and financial savings. And remember, you just need to find one of them to benefit!
Few of us have the time to stay up to date on our research given the sheer number of papers published every day.
However, given the lack of awareness in that area, just a Google search (or ChatGPT research) and reading the abstract will do a lot.
As we discussed, use AI with care. However, nowadays ChatGPT and other LLMs can save you a lot of time and help uncover sources that are hard to find. Also, don’t underestimate how specific you can be in your prompts - you can specify settings, preferences, and other requirements.
Let me give you an example:
In an institute where I recently worked as an advisor, one PhD student had leveraged the literature in an astounding manner.
What she came across was called the dropping technique:
In essence, instead of streaking bacteria across an entire plate, one would pipette dilutions directly onto the plate using a pipette.
At the top, traditional streaking is shown. Below, Chen et al. outline their approach. However, in 2015, Thomas et al. described single plate–serial dilution spotting as a similar yet more effective approach.
The effect: sensitivity remains basically unchanged (whether bacteria or fungi), with the added advantage of being able to easily test several dilutions and, importantly, improved bacterial viability.
This last is an essential point, because we often simply copy methods without truly diving deep into their nuances.
As Thomas et al. could show, when using a plastic spreader, both overly long spreading and overly long drying times can noticeably reduce bacterial viability.
Using the drop method not only removes these risks, it also has a very practical advantage. Chen et al. found that they saved about 80% of handling time.
And of course, we save resources.
Since fewer plates are used, no spreaders are needed, and tips for a multipipette are used instead, the person I referred to and her student saved more than 85% of plastic waste.
The takeaway: less time, better viability, and all of that with fewer resources. Notwithstanding that old papers are often judged outdated and literature research takes time upfront, it is often worth it.
Don’t Overlook Simplicity
However, there is a plethora of options that don’t require us to spend much time at all.
As in many laboratories, microscopy was also used in the lab I was consulting.
To optimize, it doesn’t matter what you have under your microscope - tissue, bacteria, fungi, or materials. Of note, this image, a cross-section of the colon of a germ-free mouse colonized with Candida albicans (a pathogenic fungus), won second prize in the Microverse Photo Contest. You can read more about it here.
In contrast to literature reviews and remarkable method optimization, here we find an opportunity so simple that few notice it:
Put more samples on one slide.
That’s all : )
Of note, in this case, samples were prevented from drying out using a sealant applied around them (Cytoseal, nail polish, etc.). In theory, laying samples closer could mean less sealant use if one surrounds all samples. If parafilm is used - less parafilm. PS for delicate samples in a staining chamber with reservoirs I found not parafilm lead to better tissue quality.
There are several advantages to this approach:
First, we reduce the overall number of slides.
This matters because no glass slide used in the lab is ever recycled. They are autoclaved (which requires energy) and then transported to landfills (for labs in the EU often in Africa or Asia), where they rest for decades to come, leaching chemicals into the surrounding environment.
This also leads to cost savings. While glass slides are not cheap, saving antibodies makes a real difference.
When we put samples closer together, we need less overall fluid to stain them, which means we need less expensive antibody overall.
However, since most of us will know issues with lot-to-lot variation in non-standard antibodies, by using less, we can process a larger number of samples without introducing this confounding factor.
As described previously, another advantage is that we can include more samples and controls on a single slide.
This allows us to assess not only biological variation but also staining variation, since all samples are stained on the very same slide.
Finally, we save a lot of time.
When we analyze samples, especially when using automatic scanning microscopes, we don’t need to exchange slides as often and can simply keep working, or let the microscope scan a larger area without interruption (especially important for overnight scans).
Moving Boldly, but Well-Controlled
But when do we know when we go too far?
Premeditation and controls are my key words.
The point is that many of us are inherently anxious about contamination or fragile procedures. While this concern is justified in some cases, in many others it is not.
Think back to our lesson about closing biosafety cabinets to avoid having them run overnight. Often, our concerns are excessive - therefore, trials and controls are important.
For example, a recently published paper showed that for multilocus genotype assessments, researchers were able to reuse detection 96-well plates used to assess PCR products.
We would probably not have assumed that any kind of plate could be reused in a PCR-related setting, given the high amount of genetic material involved.
This graphic comes from Berthelsen et al., outlining their experimental strategy. The paper relies heavily on statistical analyses, which can make some of the numbers difficult to follow; however, the final results and their interpretation are straightforward. And yes, an important factor to make our laboratories greener is to test ideas that may not initially seem feasible.
They achieved this by simply washing the plates with water and soap, without any harsh or time-consuming treatments.
Importantly, they tested this approach thoroughly and identified clear limitations.
For instance, the PCR plates in which the PCR reaction itself was run could not be reused, as the error rate increased.
> Therefore, when you have an idea, test it.
Of course, this requires some initial investment of resources, but the long-term savings—and the fact that others can learn from your results—often outweigh those upfront costs.
Applying The Knowledge
First, take the time to search for relevant literature, including older studies.
One year after proposing the drop technique, Sharpe and his group published another paper introducing an apparatus for dispensing drops. It featured a foot-operated diluter/dispenser and a projection viewer. Interestingly, the authors also suggested using thick-walled plastic drinking straws instead of glass pipettes to avoid clogging. Of course, over time, other techniques have been developed, such as track plating or single-plate serial dilution spotting.
Apart from time, resource, and quality improvements, this literature often discusses sources of variation we no longer think about.
In other words, it may help you better understand variability in your data.
Barbosa et al., for example, even compared three different scientists, showing how much personal handling alone can make a difference.
PS: How I analyze these papers: To make it work, you need to be efficient. Google searching often feels time-intensive although it rarely is (you can also ask ChatGPT to optimize your Google search using what are called operators). As you can see in the picture above, I normally open 10–20 tabs at a time. In my experience, more than that will eventually make you lose track. Then I read the abstract and decide whether a paper is worth reading; otherwise, I close the tab. Once I’ve selected my papers, I screen the methods first, then the results (the discussion if things are complicated, and the introduction if I need more papers). Finally, I copy-paste the title, DOI, or link, together with a 2–3 sentence summary (or parts of the abstract). You don’t need to be exhaustive upfront. I do this in word as I find it faster than citation software. I also use Firefox as it allows me to use control + mouse marking several text pieces. Information per minute is the key metric in these situations.
Still, even if you shouldn’t find options for your experiments, simple changes are almost always hiding in our protocols.
Luckily, the more trivial a change is, the safer and easier it usually is to implement.
Turning off equipment, changing the holding temperature of PCR cyclers, or simply using smaller consumables generally does not increase risk, yet can save substantial amounts of energy and plastic. Some of these unique and simple practices are discussed in this lesson. However, the real challenge is developing sensitivity and an open eye for these opportunities.
The key to identify such options is not to be held back by assumptions or fears.
Still, in some cases, even if error rates increase slightly or some spillover occurs, this may be acceptable when working with process controls or qualitative data.
But no, not everything can be optimized - and that’s fine.
How We Feel Today
References
Sharpe, A.N., et al., 1971. A rapid, inexpensive bacterial count technique using agar droplets. Journal of Applied Bacteriology, 34(2), pp.435–440. doi:10.1111/j.1365-2672.1971.tb02303.x.
Chen, C.Y., et al., 2003. A 6 × 6 drop plate method for simultaneous colony counting and MPN enumeration of Campylobacter jejuni, Listeria monocytogenes, and Escherichia coli. Journal of Microbiological Methods, 55(2), pp.475–479. doi:10.1016/S0167-7012(03)00194-5.
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.
Thomas, P., et al., 2012. Nonrecovery of varying proportions of viable bacteria during spread plating governed by the extent of spreader usage and proposal for an alternate spotting-spreading approach to maximize the CFU. Journal of Applied Microbiology, 113(2), pp.339–350. doi:10.1111/j.1365-2672.2012.05327.x.
Hoben, H.J., et al., 1982. Comparison of the pour, spread, and drop plate methods for enumeration of Rhizobium spp. in inoculants made from presterilized peat. Applied and Environmental Microbiology, 44(5), pp.1246–1247. doi:10.1128/aem.44.5.1246-1247.1982.
Barbosa, H.R., et al., 1995. Counting of viable cluster-forming and non-cluster-forming bacteria: a comparison between the drop and the spread methods. Journal of Microbiological Methods, 22(1), pp.39–50. doi:10.1016/0167-7012(94)00062-C.
Hedges, A.J., et al., 1978. Comparison of the precision obtained in counting viable bacteria by the spiral plate maker, the droplette and the Miles & Misra methods. Journal of Applied Bacteriology, 45(1), pp.57–65. doi:10.1111/j.1365-2672.1978.tb04198.x.
Berthelsen, A.L., et al., 2025. Sustainability in the laboratory: evaluating the reusability of microtitre plates for PCR and fragment detection. Royal Society Open Science, 12(5), 242226. doi:10.1098/rsos.242226.
Sharpe, A.N., et al., 1972. Technique and apparatus for rapid and inexpensive enumeration of bacteria. Applied Microbiology, 24(1), pp.4–7. doi:10.1128/am.24.1.4-7.1972.
Jett, B.D., et al., 1997. Simplified agar plate method for quantifying viable bacteria. BioTechniques, 23(4), pp.648–650. doi:10.2144/97234bm22.
Boukouvalas, D.T., et al., 2019. Automatic segmentation method for CFU counting in single plate-serial dilution. Chemometrics and Intelligent Laboratory Systems, 195, 103889. doi:10.1016/j.chemolab.2019.103889.
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Edited by Patrick Penndorf Connection@ReAdvance.com Lutherstraße 159, 07743, Jena, Thuringia, Germany Data Protection & Impressum If you think we do a bad job: Unsubscribe
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