Checklist for Experimental Design

October 25, 2021 at 9:42 am | | everyday science, scientific integrity

One of the worst feelings as a scientist is to realize that you performed n-1 of the controls you needed to really answer the question you’re trying to solve.

My #1 top recommendation when starting a new set of experiments is to write down a step-by-step plan. Include what controls you plan to do and how many replicates. Exact details like concentrations are less important than the overarching experimental plan. Ask for feedback from your PI, but also from a collaborator or labmate.

Here are some things I consider when starting an experiment. Feel free to leave comments about things I’ve missed.

Randomization and Independence

  • Consider what “independent” means in the context of your experiment.
  • Calculate p-value using independent samples (biological replicates), not the total number of cells, unless you use advanced hierarchical analyses (see: previous blog post and Lord et al.:
  • Pairing or “blocking” samples (e.g. splitting one aliquot into treatment and control) helps reduce confounding parameters, such as passage number, location in incubator, cell confluence, etc.

Power and Statistics

  • Statistical power is to the ability to distinguish small but real differences between conditions/populations.
  • Increasing the number of independent experimental rounds (AKA “biological replicates”) typically has a much larger influence on power than the cells or number of measurements per sample (see: Blainey et al.:
  • If the power of an assay is known, you can calculate the number of samples required to be confident you will be able to observe an effect.
  • Consider preregistering
    • Planning the experimental and analysis design before starting the experiment substantially reduces the chances false positives.
    • Although formal preregistration is not typically required for cell biology studies, simply writing the plan down for yourself in your notebook is far better than winging it as you go.
    • Plan the number of biological replicates before running statistical analysis. If you instead check if a signal is “significant” between rounds of experiment and stop when p < 0.05, you’re all but guaranteed to find a false result.
    • Similarly, don’t try several tests until you stumble upon one that gives a “significant” p-value.


  • “Blocking” means to subdivide samples into similar units and running those sets together. For example, splitting a single flask of cells into two, one treatment and one control.
  • Blocking can help reveal effects even when experimental error or sample-to-sample variability is large.
  • Blocked analyses include paired t-test or normalizing the treatment signal to the control within each block.


  • Sledgehammer controls
    • These controls are virtually guaranteed to give you zero or full signal, and are a nice simple test of the system.
    • Examples include wild type cells, treating with DMSO, imaging autofluorescence of cells not expressing GFP, etc.
  • Subtle controls
    • These controls are more subtle than the strong controls, and might reveal some unexpected failures.
    • Examples include: using secondary antibody only, checking for bleed-through and crosstalk between fluorescence channels, and using scrambled siRNA.
  • Positive & negative controls
    • The “assay window” is a measure of the range between the maximum expected signal (positive control) and the baseline (negative control).
    • A quantitative measure of the assay window could be a standardized effect size, like Cohen’s d, calculated with multiple positive and negative controls.
    • In practice, few cell biologist perform multiple control runs before an experiment. So a qualitative estimate of the assay window should be considered using the expected signal and expected variability sample to sample. In other words, consider carefully if an experiment can possibly work
  • Concurrent vs historical controls
    • Running positive and/or negative control in the same day’s experimental run as the samples that receive real treatment helps eliminate additional variability.
  • Internal controls
    • “Internal” controls are cells within the same sample that randomly receive treatment or control. For example, during a transient transfection, only a portion of the cells may actually end up expressing, while those that aren’t can act as a negative control.
    • Because cells with the same sample experience the same perturbations (such as position in incubator, passage number, media age) except for the treatment of interest, internal controls can remove many spurious variables and make analysis more straightforward.


  • Blinding acquisition
    • Often as simple as having a labmate put tape over labels on your samples and label them with a dummy index. Confirm that your coworker actually writes down the key, so later you can decode the dummy index back to the true sample information.
    • In cases where true blinding is impractical, the selection of cells to image/collect should be randomized (e.g. set random coordinates for the microscope stage) or otherwise designed to avoid bias (e.g. selecting cells using transmitted light or DAPI).
  • Blinding analysis
    • Ideally, image analysis would be done entirely by algorithms and computers, but often the most practical and effective approach is old-fashioned human eye.
    • Ensuring your manual analysis isn’t biased is usually as simple as scrambling filenames. For microscopy data, Steve Royle‘s macro, which works well:
    • I would highly recommend copying all the data to a new folder before you perform any filename changes. Then test the program forward and backwards to confirm everything works as expected. Maybe perform analysis in batches, so in case something goes awry, you don’t lose all that work.


Great primer on experimental design and analysis, especially for the cell biologist or microscopist: Stephen Royle, “The Digital Cell: Cell Biology as a Data Science”–eqskudatarq=1282

Advanced, detailed (but easily digestible) book on experimental design and statistics: Stanley Lazic, “Experimental Design for Laboratory Biologists”

I like this very useful and easy-to-follow stats book: Whitlock & Schluter, “The Analysis of Biological Data”

Alex Reinhart, “Statistics Done Wrong”

SuperPlots: Communicating reproducibility and variability in cell biology. (HTMLPDF)
Lord, S. J.; Velle, K. B.; Mullins, R. D.; Fritz-Laylin, L. K. J. Cell Biol.2020219(6), e202001064.

update on Nikon objective immersion oils

August 30, 2018 at 8:41 am | | everyday science, hardware, review

A few years ago, I compared different immersion oils. I concluded that Nikon A was the best for routine fluorescence because: (A) it had low autofluorescence, (B) it didn’t smell, (C) it was low viscosity, and (D) the small plastic dropper bottles allowed for easy and clean application.

Unfortunately, my two favorites, Nikon A and NF, were both discontinued. The oil Nikon replaced these with is called F. But I don’t love this oil for a few reasons. First, it’s fairly stinky. Not offensive, but I still don’t want my microscopes smelling if I can help it. Second, I’ve heard complaints from others that Nikon F can have microbubbles (or maybe crystals?) in the oil, making image quality worse. Finally, dried F oil hardens over time, and can form a lacquer unless it is cleaned off surfaces very well. That said, F does have very low fluorescence, so that’s a good thing.

I explored some alternatives. Cargille LDF has the same optical properties as Nikon F (index of refraction = 1.518 and Abbe Ve = 41). But LDF smells terrible. I refuse to have my microscope room smell like that! Cargille HF doesn’t smell and has similar optical properties, but HF is autofluorescent at 488 and 405 nm excitation, so it adds significant background and isn’t usable for sensitive imaging.

At the recommendation of Kari in the UCSF microscopy core (and Caroline Mrejen at Olympus), I tried Olympus Type F, which also has an index of refraction of 1.518 and an Abbe number of 40.8, which is compatible with Nikon. The Olympus oil had very low autofluorescence, on par with Nikon A, NF, and F. (I also tested low-fluorescence oils Leica F and Zeiss 518F, but their dispersion numbers are higher (Ve = 45-46), which can cause chromatic aberration and may interfere with Perfect Focus.)

I used to love the low viscosity of Nikon A (150 cSt), because it allowed faster settling after the stage moved and was less likely to cause Perfect Focus cycling due to mechanical coupling to thin or light samples, plus it was easier to apply and clean. Nikon NF was higher viscosity (800 cSt). Olympus F is higher than Nikon A (450 cSt), but acceptable.

Finally, Olympus F comes is an easy to use applicator bottle: instead of a glass rod that can drip down the side of the vial if you’re not careful, the Olympus F is in a plastic bottle with a dropper. It’s not quite as nice as the 8 cc dropper bottles that Nikon A used to come in, and I don’t love the capping mechanism on the Olympus F, but I’ll survive.

I plan to finish up our last bottle of Nikon A, then switch over to Olympus F. We also have a couple bottles of Nikon NF remaining, which I will save for 37C work (the higher viscosity is useful at higher temperatures).


Some people claim that type A was simply renamed type N. I don’t think that’s true. First of all, I couldn’t get Perfect Focus on our Ti2 to work with Nikon type N oil. Second, the autofluorescence of Nikon type N (right) was way higher than Olympus type F (left) or the old Nikon type A, at least at 405 and 488 nm:

So I’ll stick with Olympus type F. :)


Here are some example images. These are excited with 640 (red), 561 (red), 488 (green), and 405 nm (blue) and the display ranges are the same for each sample. (The dots are single fluorophores on the glass.) You can see that Cargille HF is slightly more autofluorescent (especially at 405 nm) than either the old Nikon A or Olympus type F. This matches what Cargille states for HF: “Slightly more fluorescent than Type LDF.”

always have plastic sheeting in lab

October 24, 2016 at 10:46 am | | everyday science, hardware, stupid technology

I learned during my PhD that you should always have plastic sheeting in lab, because it might just save your equipment when/if a water leak happens. It saved one of our scopes recently, although I wasn’t fast enough to prevent some water damage on an expensive camera. :(

2016-09-02 14.27.09

2016-09-02 14.17.25

For less than $5, you can get some rolls of the stuff. If you want larger and thicker sheets (like in the photos), I recommend this stuff.

dissertation acknowledgments

March 19, 2015 at 2:27 pm | | everyday science, grad life, history

Paul and ChemJobber posted about acknowledgements in theses and dissertations. Paul has a nice one here. It made me reread mine:


Most importantly, I thank my advisor, W.E. Moerner. It is difficult to explain how wonderful it has been to study under him. W.E. is a real scientist’s scientist: he fundamentally cares about good science and presenting results in a clear and honest manner. He always impressed me with his understanding of sciences outside his field and his scholarship, as I doubt that there is any paper I have read that he has not. W.E. always knows where some obscure piece of equipment is in the lab, and what type of power cable it requires. W.E.’s humor and generosity have been invaluable during my time in his lab, not to mention his scientific guidance. I could not have asked for a better Ph.D. advisor.

I joined the Moerner lab because W.E. seemed to run a fun and exciting research program, and I have not been disappointed. Other members of the Moerner Lab have been instrumental in my education and research. Kallie Willets mentored me when I first arrived at Stanford. Kallie was fun to work with and I am very grateful for the time and energy she dedicated to helping me get a solid footing in the lab by teaching me the right way to do things (and clean up afterwards).

After Kallie graduated, it was entertaining (to say the least) to get to know my officemate Dave Fromm. Dave was always willing to discuss problems I was facing in my experiments, and often suggested perfect solutions. (He was also always willing to discuss his adventures and funny stuff he found on the internet.) Dave and Jim Schuck regularly played darts over my head … literally. In general, this was entertaining and helpful to my overall spirit, and I appreciate the fun times with Jim and Dave. In those early years, I also enjoyed the company of (and scientific input from) Nick Conley, Anika Kinkhabwala, Adam Cohen, Stefanie Nishimura, Jaesuk Hwang, Kit Werley, So Yeon Kim, Andrea Kurtz, Marcelle Koenig, and Jian Cui.

In the later years of my tenure in the Moerner Lab, I have benefited from another batch of amazing people. Nick is one of the most motivating collaborators I have had the pleasure of working with; he is always excited about results, and his mind wanders to great places (not to mention that his skills as an organic chemist were very helpful to me)! I also had the opportunity to work with Hsiao-lu Lee, who was always generous with her time and expertise in cell culture. I am thankful to have those two wonderful coauthors. Alex Fürstenberg has been a fun (and very tolerant) officemate, and is always a great person to ask about anything photophysical. Mike Thompson is hard working and smart, but most importantly he laughs at more than 83% of my jokes. Julie Biteen is opinionated and usually right, and has been fun to bounce ideas off. All the other members of the Moerner Lab (Shigeki, Randy, Majid, Steve, Jianwei, Whitney, Lana, Yan, Sam B, Quan, Matt, etc.) are exceptional people and have made Stanford a wonderful place.

Marissa Lee started joined the lab in 2008, joining my project. I have enjoyed mentoring her and passing on as much as possible of what Kallie, Dave, Jim, Stefanie, Nick, Hsiao-lu, So Yeon, Jaesuk, Adam, and Anika taught me over the years. I wish her luck in her time at Stanford. Several summer students worked with me to get a taste of research. I thank Jennifer Alyono, Daniel Lau, Nathan Hobbs, and John Servanda for their help taking spectra.

Of course, I must also acknowledge Bob Twieg and his students at Kent State University. As a physical chemist, there is nothing better than an excellent collaboration with a group of top-notch synthetic chemists. W.E. and Bob have worked together since their IBM days in the 1980s and 1990s, and I had the fortune to benefit immensely from that bond between labs. Nearly every compound mentioned in this Dissertation was synthesized by the Twieg lab, and the back-and-forth (or push– pull?) design process between the labs should serve as an example to what all collaborations should strive for. Bob’s students have made great compounds over the years, and I thank all of them for being super collaborators: Meng, Hui, Zhikuan, Na, Reichel, Ryan, Alex, and Jarrod.

Friends have made grad school a blast. I met the Moilanens immediately, and enjoyed marathon training and adventures with David and Hailey. Ben Spry was a great help studying for placement exams, and I enjoyed driving to San Jose with Ben so he could buy a Camaro. William Childs and Charles McCrory—after I finally decided to like them—were indispensible: grad school will be filled with fond memories of coffee, lunch, and arguments because of Wm and Charles. So many other friends made my time at Stanford wonderful: Nichole, Kate, Alicia, Jen, Drew, Ashley, John, Zalatan, Chad, Matt, Griffin, Kendall, Daniel, Adrienne, Adam, Avisek, Eric, Ethan, Kevin, Emily, Ken, Dan, Scott, and everyone else! It has been fun having Jordan and Maria in California, and so many other non-Stanford friends that I cannot possibly name them all. I have had positive interactions with several faculty members, and I thank Bob Waymouth, Chris Chidsey, Dick Zare, Bianxiao Cui, Steve Boxer, Justin DuBois, Vijay Pande, Bob Pecora, and Ed Solomon. I also must recognize the members of the Stanford staff who contributed to my work and enjoyment, namely: Roger Kuhn, Todd Eberspacher, Brian Palermo, Patricia Dwyer, Grace Baysinger, Steve Lynch, and all the Conways—Marc, Daragh, and Mariette.

I feel that I must also acknowledge those in my past who influenced me and led me down the path of science. My earliest memories of enjoying the natural world were at Audubon’s Mast Landing Camp, playing and learning about nature with Aaron and Ira and Matt. In the third grade, Mrs. Solari recognized and encouraged my inclination toward science, as have many teachers since. I thank Dr. Root, who mentored me for my 7th-grade science fair project; Mr. Plummer for dealing with 8th graders; Mr. Glick for the astronomy and recycling clubs and supporting me throughout high school; Mr. Herrick, for being the best physics teacher I never had; Mr. Gauger for insisting that Heisenberg’s uncertainty principle can explain why things still jiggle at zero Kelvin; John Anderson for arguing with me; Don Cass for teaching my first college chemistry class and making it so exciting; Tony Planchart for teaching biochemistry in a way that convinced me to be a chemistry major; Helen Hess for fun classes biology and biomechanics; Michael Rubinstein for his entertaining exploration of polymer physics; Royce Murray for teaching analytical chemistry; Max Berkowitz for stat mech classes; and Charles Schroeder, Eric Shaqfeh, and Steve Chu for a great summer research experience. I should offer a special bit of gratitude to Sergei Sheiko, whose lab I worked in as an undergrad, and who helped make my time at UNC spectacular.

This Dissertation is dedicated to my family: the Lords, the Cyrs, and the Hearns. My parents have always encouraged my interests, without pushing me too hard. I wouldn’t be half the person I am without their support. My brother Jackson has been a life-long companion, so I was very pleased when he moved to California and we could play together like when we were growing up. My grandparents Lord funded my education, which I greatly appreciate. I probably get some of my curiosity from my pépère Cyr. My first year at Stanford, I met Brenna Hearn and married her a few years later. She has made my life wonderful, and I thank her for her support throughout grad school. I cannot thank Brenna enough for her companionship, so I’ll stop there.

Looking back at this, I wish that I had made it 50 times longer and cut out the rest of the dissertation.

how a biochemist siphons

January 30, 2015 at 10:56 am | | everyday science

2015-01-27 18.46.38

comparing Nikon immersion oils

August 27, 2014 at 2:53 pm | | everyday science, review

UPDATE: update on Nikon objective immersion oils

I typically use Nikon type NF immersion oil. But I hate the dropper that it comes in, and I’ve recently been having trouble with the oil crystallizing, especially if I aliquot it to smaller dropper vials. So I decided to compare the different oil types available, namely Nikon A (not to be confused with Cargille A), Cargille B, Cargille 37, and Nikon NF. (Type 37 is sometimes called type B 37.) Note that types B and 37 are actually Cargille part numbers 16484 and 16237, respectively.

A B 37 NF montage

See full slide deck here.

My conclusion: Use Nikon A for routine imaging (the dropper is much easier to use and it’s less stinky than NF). For samples at 37 C or single-molecule imaging, use type NF.

UPDATE: Unfortunately, Nikon has discontinued their type A and NF oils. Back to the drawing board. I will update with what oil I chose to use in the future

home-made plasma cleaner

July 23, 2014 at 12:52 pm | | everyday science, hardware, science@home, stupid technology

I really want a plasma cleaner, for cleaning coverslips and activating glass for PDMS bonding, but they cost thousands of dollars. I thought that was a lot of money for a glorified microwave. So I made my own.

Drill a few holes in glass:

2014-03-10 11.16.58


Make a PDMS seal (thanks Kate):

2014-07-16 16.11.57

Glue the chamber:

2014-03-10 12.29.04

We’re ready to go!

2014-07-17 13.19.41

Fill the chamber with argon, evacuate it, turn on the microwave oven, and … voila! … a plasma:

2014-07-17 12.54.18

2014-07-17 12.56.04

Below are slides before and after (right) plasma treatment. You can see the contact angle of water is dramatically reduced.

2014-07-17 13.14.40

It works!

Well, not really. I found that the plasma really only stays lit with argon. When I flow air in, it extinguishes, but also burns some of the rubber hoses. That adds more dirt to my slides than I want.

Conclusion: don’t do this at home. :)

(Well, that might be a little harsh. It does work well to bond PDMS to glass. And I’ll try a longer etch sometime to see if it will ever clean the coverslips.)

thorlabs lab snacks boxes for arduino enclosure

July 3, 2014 at 7:38 pm | | everyday science, hardware

On the topic of hardware syncing, I figured I should boast about my very fancy Arduino enclosure. I used a Thorlabs Lab Snacks box (one of the Great Boxes of Science):

2014-06-11 18.17.49 2014-06-11 18.17.33

2014-06-11 18.17.26

Of course, Nico makes beautiful laser-cut boxes for his Arduino, and Kurt has a nice 3D-printed box. But I think I’ll stick to this reduce/reuse/recycle approach. :)

UPDATE: I guess I’m not the only one. Labrigger posted a similar pic!

UPDATE 2: I made a bigger one to fit two Arduinos:

2014-08-07 14.07.19 2014-08-07 14.10.30

ActiveView PDF

April 10, 2013 at 10:38 am | | everyday science, literature, news

Does anyone else love ACS’s ActiveView PDF viewer for reading PDFs and seeing reference? And Nature’s ReadCube, too. Great stuff.

Of course, after I scan the ActiveView, I still download the old-fashioned PDF and use Papers (or Mendeley) to read and manage my library.

google reader alternatives

April 3, 2013 at 8:12 am | | everyday science, literature, science community, software

Now that Google Reader is going the way of the dodo Google Gears, how am I going to keep up with the literature?!? I read RSS feeds of many journal table of contents, because it’s one of the best ways to keep up with all the articles out there (and see the awesome TOC art). So what am I to do?

There are many RSS readers out there (one of my favorites was Feeddler for iOS), but the real problem is syncing! Google servers took care of all the syncing when I read RSS feeds on my phone and then want to continue reading at home on my computer. The RSS readers out there are simply pretty faces on top of Google Reader’s guts.

But now those RSS programs are scrambling to build their own syncing databases. Feedly, one of the frontrunners to come out of the Google Reader retirement, claims that their project Normandy will take care of everything seamlessly. Reeder, another very popular reader, also claims that syncing will continue, probably using Feedbin. Feeddler also says they’re not going away, but with no details. After July 1, we’ll see how many of these programs actually work!

So what am I doing? I’ve tried Feedly and really like how pretty it is and easy it is to use. The real problem with Feedly is that its designed for beauty, not necessarily utility. For instance look how pretty it displays on my iPad:


But note that its hard to distinguish the journal from the authors and the abstract. And it doesn’t show the full TOC image. Feedly might be faster (you can swipe to move to the next articles), but you may not get as much full information in your brain and might miss articles that might actually interest you.

Here’s Reeder, which displays the title, journal, authors, and TOC art all differently, making it easy to quickly scan each  article:



And Feeddler:


I love that Feeddler lets me put the navigation arrow on the bottom right or left, and that it displays a lot of information in nice formatting for each entry. That way, I can quickly flip through many articles and get the full information. The major problem is that it doesn’t have a Mac or PC version, so you’ll be stuck on your phone.

I think I’ll drop Feeddler and keep demoing Reedler and Feedly until July 1 rolls around.

sam’s three most important safety rules

October 12, 2011 at 4:09 pm | | everyday science, lab safety

Top three safety rules, especially for new students:

  1. If you’re unsure about any safety issue, ask someone!
  2. Wear safety glasses when freezing things.
  3. Wear a face shield when piranha etching.

An addendum rule is to not work sloppily in general. Or when you’re very tired.

Of course, there are many other important rules. But these are my favs.

Which antibiotic resistance gene does my plasmid have?

February 8, 2011 at 2:28 pm | | cool results, everyday science

Few experiments in science are conclusive.  So, it is very exciting when an experiment provides completely unambiguous results.  Today that happened.

Molecular biologists use bacteria–specifically strains of E. coli that don’t make people sick (i.e., non-pathogenic)–to make lots of copies of circular DNA, called plasmids; one can think of E. coli as a photocopier for plasmids.  Bacteria love to take up plasmids, copy them, and express them to make proteins with unique functions, and it is this ability that makes them so evolutionarily successful.  Bacteria can transfer plasmids containing antibiotic resistance genes to each other, for example, and in this manner, can become “superbugs.”  Staph is one pathogen that has done this so successfully in hospitals that we will soon run out of antibiotics to treat it.

Interestingly, molecular biologists give E. coli antibiotic resistance genes on purpose. It’s not because we’re bioterrorists.  Rather, we want to be able to SELECT for those bacteria that take up our plasmid of interest and copy it.  So we make sure that the plasmid DNA that we give the E. coli to copy also has a gene that codes for a protein that confers antibiotic resistance; ampicillin resistance is the most common.  We take this plasmid, mix it with the E. coli, and warm the E. coli slightly.  This creates little pores in the E. coli that allow the plasmid DNA to pass through.  The process–called heat transformation–is not very efficient, and most of the E. coli don’t take up any plasmids.  We don’t want to grow these E. coli because they are useless to us; we only want to grow the ones that took up our plasmid.  So, we add some ampicillin, and only those E. coli that took up our plasmid DNA can survive.  As a result, we end up with a bacterial culture that is loaded with our “photocopied” DNA.  The resulting plasmid DNA can be used for many things; for example, DNA that codes for insulin can be copied in E. coli, purified, and then put into mammalian cells to cause them to make insulin, which can be harvested and given to diabetic patients.

I recently wanted to use E. coli to “photocopy” some plasmid DNA that I got from another researcher, but the researcher wasn’t sure which antibiotic resistance gene for selection was employed in the plasmid (not good record keeping).  He thought it was either ampicillin or kanamycin.  So, I transformed E. coli with the plasmid and tried growing the bacteria in both ampicillin and kanamycin.  Take a look at the picture below.  It’s pretty obvious that the bacteria survived only in the kanamycin (yellow cloudy suspension on the right), indicating that the plasmid coded for kanamycin resistance.  The bacteria in the ampicillin died quickly and did not grow.

now that’s pure!

December 15, 2010 at 11:54 am | | everyday science, great finds

That’s very pure EDTA:

100.06% pure!

dangerous chemists need to be fired

August 25, 2010 at 2:32 pm | | everyday science, lab safety


I don’t mean that if someone makes one mistake they’re gone, but if they consistently put themselves and others in danger, they should be fired. Or do theory.

Dangerous chemists are the ones who don’t know how to perform labwork safely, but think they do or refuse to ask for guidance. They are the ones that everyone else in lab knows is unsafe. They make the same mistakes over and over. They regularly work in lab alone. They don’t update their labmates about the dangerous compounds or reactions they are using.

Dangerous chemists are inconsiderate, put others at risk, and should be fired.

I started writing this a while back because I’d heard stories from friends about a chemist. His labmates are scared of his experiments because he was reckless, ignorant, and didn’t talk to people about what he is doing. He performed dangerous reactions on the work bench instead of the hood.

For instance, his labmates noticed that he was holding his breath while in lab, making adjustments to some reaction. When asked why he was holding his breath, he answered that the reaction produced dangerous fumes. So, instead of properly venting the reaction, discussing the reaction with his lab/PI, or warning his labmates, he just held his breath.

And yet this chemist was allowed to continue working in lab, even after many complaints to his PI and others in authority. He should have just been fired; I’m sure there’d be someone capable eager to take his spot!

I wasn’t going to post this, because it there were some funnier things to blog. But now the story of Preston Brown blowing off his fingers after grinding up 10 g of very explosive hydrazine. (The returned ChemBark also blogged this!) Now, I feel sad for Brown: he did not deserve to be injured, even if he was being reckless. But I also think that he should have been fired long before this accident occurred. It sounds like his labmates knew he was dangerous.

If there’s someone in your lab who you think is dangerous (who, if he or she blew up the lab, your first reaction would be, I saw that coming), do the following:

  1. First talk to him/her. Voice your concerns and offer to help train him/her in proper technique.
  2. Talk to your PI. State clearly that you are concerned for your personal safety in lab because of your labmate’s dangerous behavior. Make sure the lab has an SOP for every dangerous procedure in lab. And make sure the SOP is enforced.
  3. Talk to the authorities. If your PI refuses to make the situation safe, go to the department safety coordinator or the EH&S.
  4. Refuse to work around dangerous chemists. It’s not worth putting your life at risk. You’d have ground for a lawsuit if they fire you for refusing to work in an unsafe environment. (Hell, you may not even be covered by worker’s comp if the idiot hurts you!) Stand up for your rights: grad school is not a sweatshop.
  5. Document. Save emails and sent paper letters, just in case you need to sue. ;)

That said, I suspect that most things never have to go past step 1. I think most dangerous things done is lab are mistakes or lack of understanding of the correct protocols. Rarely, someone repeatedly ignores protocols and their PI’s instructions to intentionally perform dangerous experiments. But those rare instances is what I’m talking about. Those fools need to be fired.

I do want to note, Brown’s story is not final. Maybe it’ll come out that he was not as reckless as it seems from the C&E News article. In fact, it’s entirely possible, that he just wasn’t trained well enough. My point is that it is his PI’s responsibility to train him in safety, and fire him when he refused to be safe!

And I want to say again that he did not deserve to be injured. I feel really bad for him. A quote from the investigation transcript: “OK. Thanks again for coming to the house. I know. It’s a little more hassle. … I was left handed. I’ll have to be right handed now.”

UPDATE: Chemjobber has a nice post about why the faculty members bear some responsibility in the Texas Tech case.

UPDATE 2: AGAM has a post reminding us not to be too cocky in our safety knowledge. A good point: we all should regularly be boning up on our safety train, and communicating with our colleagues about best practices. Here are links to Prudent Practices and working with azides.

Photojournalism Tour

August 12, 2010 at 1:25 am | | everyday science, grad life, lab safety

These photos are from anonymous labs:

Like working at the MMS.

A monument to Thorlabs.

Money well spent.

Laser safety.

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