Bento Lab

Bioinformatics tool review 👇

Here's a comprehensive online suite of old-school DNA and protein sequence manipulation tools called the "Sequence Manipulation Suite". It's been in constant use since 2000, probably because it's so useful and accessible for everyone from beginners to teachers to researchers!

The Sequence Manipulation Suite, by Paul Stothard, can be used in a web browser online or offline (you can download it for offline use). It runs on Javascript so you'll need this enabled on your browser for it to work.

It contains a large number of individual “simple” operations that involve copying your sequence (or text) into a text box, selecting parameters or adding additional sequence strings (for example primers), and then clicking to submit the operation.

It has more manipulation functions than can easily be listed here, but useful manipulations from a perspective (for example) include:

⭐ Reverse complementing DNA sequences
⭐ Converting GenBank sequences to FASTA files
⭐ Mapping primers to a sequence
⭐ Simple pairwise alignment
⭐ Fuzzy searching within a DNA sequence
⭐ Mapping restriction enzyme cut points

And from a quick glance, some uses in recently published articles include:

🔍 Identifying amino acid codons
🔍 Calculating molecular weights of proteins
🔍 Calculating percentage identity and amino acid similarity between sequences
🔍 Visualising similarities between sequences in multiple sequence alignments

The original article describing it has 1818 citations (according to Google Scholar) across 23 years; it’s been recently updated to make it run more smoothly; and it’s still in use today.

You can find the Sequence Manipulation Suite here: https://bioinformatics.org/sms2/

And you can read its original publication here:

Stothard, Paul. "The sequence manipulation suite: JavaScript programs for analyzing and formatting protein and DNA sequences." Biotechniques 28.6 (2000): 1102-1104. https://www.future-science.com/doi/abs/10.2144/00286ir01

Have you ever wanted to take PCR workflows off-grid? 👇

You can do this with Bento Lab (the portable PCR workstation) and a portable power station or generator — find out more here:

https://bento.bio/resources/bento-lab-advice/powering-bento-lab-in-the-field-with-a-battery/

🌳 New tool alert for those interested in drawing phylogenetic trees!

Here’s a new free and powerful tree-drawing software application called TreeViewer. Available on Windows/macOS/Linux, it lets you transform phylogenetic trees, add layers, map data to branches, align text, and much more!

Treeviewer, developed by Bianchini & Sánchez‐Baracaldo (2024), aims to be a flexible, modular and user-friendly application that can help make phylogenetic trees publication-quality without needing to do additional modifications on graphics packages such as Photoshop.

Its modular system also allows future transformations to be developed by its creators and its users.

Its features allow:

🌳 Alignment of images with branches, for example drawing taxon images next to their respective clades

🌳 Displaying alignments alongside phylogenies to show taxonomically important regions

🌳 Displaying character states next to each branch, for example taxonomically important characters or the presence and absence of specific genes

🌳 Extensive modifications using transformations, colour-coding, and additional objects

🌳 Custom scripts

The software is very powerful and has lots of different options, so it may take a little while to get used to it, but fortunately it has a detailed manual with worked examples that should get you familiar with the key functions.

TreeViewer can be downloaded from https://treeviewer.org, and you can read the accompanying article here:

Bianchini & Sánchez‐Baracaldo (2024). TreeViewer: Flexible, modular software to visualise and manipulate phylogenetic trees. Ecology and Evolution, 14(2), e10873. https://onlinelibrary.wiley.com/doi/full/10.1002/ece3.10873

📸 TreeViewer

Technique tip to remove PCR inhibitors and precipitate DNA 👇

For anyone working with PCR and "recalcitrant" plants full of PCR inhibitors, here are two simple and inexpensive DNA extraction methods that might be useful, using TE buffer, a detergent (SDS), and (unusually!) acetone to remove PCR inhibitors and precipitate DNA! Both methods are designed for fresh leaves and dried herbarium specimens by the same team.

The first method (Gouker et al. (2020)) involves a 2 hr+ acetone soak of leaf discs followed by evaporation of the acetone; manual grinding of leaf tissue with a metal rod; DNA extraction in 1x TE buffer and 1% SDS at 90 °C for 10 minutes; centrifugation and removal of the supernatant; and an acetone DNA precipitation followed by centrifugation and resuspension.

For this method, the authors found that fresh samples from a wide range of woody plants (from Acer to Quercus) gave DNA yields of between 11 ng to 1080 ng DNA, with all fresh samples giving successful amplification. However, herbarium specimens were less successful (presumably due to their age) and although DNA was extracted only half of the samples produced amplicons. Still, these were very old and difficult plant specimens...

The second method (Gouker et al. (2023)) involves homogenising two 5 mm diam. leaf discs with a FastPrep machine using a 4.5 mm ball; adding 10x TE buffer with 1% SDS and 1% PVPP; vortexing and centrifugation to precipitate solids; transferal of the supernatant to a new tube and adding acetone; centrifugation to pellet DNA, discarding the supernatant and adding nuclease-free water; vortexing then centrifuging; and saving the supernatant for DNA quantitation and downstream PCR applications. The whole process should take only 40 minutes and cost only $0.23 per sample for the extraction.

The second method is an update and improvement on the first method, but it requires the use of a FastPrep tissue homogeniser. So for some people the first method (or a hybrid of the two) may be more feasible.

The results for this method showed that the extraction method did extract more DNA from old herbarium specimens than a comparison CTAB and Qiagen Kit extraction. However, PCR success was less successful, with only 66% of samples amplifying for longer fragments and 92% for the much shorter P6 loop amplicons. This was presumably due to the continued presence of PCR inhibitors in those particular specimens or plants.

Even so, the method worked well enough that the authors suggest that it should be considered as a rapid, inexpensive option for recalcitrant plant specimens, especially given the small amount of starting tissue required.

You can read Gouker et al. (2020) and Gouker et al. (2023) here:

Gouker et al. (2020). https://bsapubs.onlinelibrary.wiley.com/doi/full/10.1002/aps3.11403
Gouker et al. (2023). https://bsapubs.onlinelibrary.wiley.com/doi/full/10.1002/aps3.11521Technique tip to remove PCR inhibitors and precipitate DNA 👇

For anyone working with PCR and "recalcitrant" plants full of PCR inhibitors, here are two simple and inexpensive DNA extraction methods that might be useful, using TE buffer, a detergent (SDS), and (unusually!) acetone to remove PCR inhibitors and precipitate DNA! Both methods are designed for fresh leaves and dried herbarium specimens by the same team.

The first method (Gouker et al. (2020)) involves a 2 hr+ acetone soak of leaf discs followed by evaporation of the acetone; manual grinding of leaf tissue with a metal rod; DNA extraction in 1x TE buffer and 1% SDS at 90 °C for 10 minutes; centrifugation and removal of the supernatant; and an acetone DNA precipitation followed by centrifugation and resuspension.

For this method, the authors found that fresh samples from a wide range of woody plants (from Acer to Quercus) gave DNA yields of between 11 ng to 1080 ng DNA, with all fresh samples giving successful amplification. However, herbarium specimens were less successful (presumably due to their age) and although DNA was extracted only half of the samples produced amplicons. Still, these were very old and difficult plant specimens...

The second method (Gouker et al. (2023)) involves homogenising two 5 mm diam. leaf discs with a FastPrep machine using a 4.5 mm ball; adding 10x TE buffer with 1% SDS and 1% PVPP; vortexing and centrifugation to precipitate solids; transferal of the supernatant to a new tube and adding acetone; centrifugation to pellet DNA, discarding the supernatant and adding nuclease-free water; vortexing then centrifuging; and saving the supernatant for DNA quantitation and downstream PCR applications. The whole process should take only 40 minutes and cost only $0.23 per sample for the extraction.

The second method is an update and improvement on the first method, but it requires the use of a FastPrep tissue homogeniser. So for some people the first method (or a hybrid of the two) may be more feasible.

The results for this method showed that the extraction method did extract more DNA from old herbarium specimens than a comparison CTAB and Qiagen Kit extraction. However, PCR success was less successful, with only 66% of samples amplifying for longer fragments and 92% for the much shorter P6 loop amplicons. This was presumably due to the continued presence of PCR inhibitors in those particular specimens or plants.

Even so, the method worked well enough that the authors suggest that it should be considered as a rapid, inexpensive option for recalcitrant plant specimens, especially given the small amount of starting tissue required.

You can read Gouker et al. (2020) and Gouker et al. (2023) here:

Gouker et al. (2020). https://bsapubs.onlinelibrary.wiley.com/doi/full/10.1002/aps3.11403
Gouker et al. (2023). https://bsapubs.onlinelibrary.wiley.com/doi/full/10.1002/aps3.11521

The first protocol has since been used successfully for extraction of boxwood and fungal DNA from infected tissue, and fungal DNA from culture, by Gouker et al. (2022):

Gouker et al. (2022). https://apsjournals.apsnet.org/doi/full/10.1094/PHYTOFR-09-21-0066-SC

It'll be interesting to see if any future modifications of these acetone-based methods can improve PCR success from the most recalcitrant plants or specimens. Maybe a quick filter paper dipstick extraction would be enough to clean up the last of the PCR inhibitors?

Please also note that even though acetone is available for household use as nail varnish remover, it is highly toxic if ingested, and volatile and flammable, so it should be treated with all due care.

Using acetone for rapid PCR‐amplifiable DNA extraction from recalcitrant woody plant taxa Premise Quick and effective DNA extraction from plants for subsequent PCR amplification is sometimes challenging when working across diverse plant taxa that may contain a variety of inhibitory compo...

2 minute journal club 📄 👇

We enjoyed reading this new approach for DNA metabarcoding microscopic protist phytoplankton, the “building blocks of aquatic food webs” using full-length 18S DNA metabarcoding.

Gaonkar et al. (2024) ask if metabarcoding marine protist phytoplankton with Oxford Nanopore MinION and full-length 18S rDNA reads (~1829 bp) could be more successful than previous methods with short read Illumina metabarcoding.

Specifically, could they improve taxonomic classifications using longer sequences and capture more of the taxonomic diversity present?

The work:

🌿 Developed a new primer set to avoid a known problem site in the 18S region, and tested these in silico.

🌿 Cultured six representatives of the major groups of marine phytoplankton, and extracted and amplified the DNA to validate the new primers.

🌿 Created mock communities using PCR products produced during their testing of their new primers; and with PCR products for the 18S V4 and V8-V9 regions for a comparison to ensure that these primers would also work for their test species.

🌿 Sampled phytoplankton from 6 sites off the coast of Texas, extracting and amplifying the DNA using the new primers and the V4 and V8-V9 primers to test their method against the Illumina MiSeq method using real-life environmental communities.

They found the duplex option of the super high accuracy basecalling model was vastly superior to the high-accuracy model under their test conditions (≤2 mismatches and ≤1 gap across ~1750 bp sequences) when sequencing the mock communities with ONT MinION.

When they sequenced the environmental samples, they found:

🔍 47% of ONT reads met their quality criteria for sequencing (≥90% similarity to existing sequences, and ≥1000 bp)

🔍 No metabarcoding primers detected all 298 genera present in the environmental samples. Full-length 18S MinION sequencing detected 250 genera (84%), while MiSeq V4 and V8-V9 identified only 226 (76%) and 213 genera (71%) respectively.

🔍 MinION full-length 18S sequencing detected almost all diatoms, ciliates, Plantae, cryptophytes, and radiolarians, but only 78% of dinoflagellates and 50% of cercozoans.

The authors suggest the detection failures could be due to primer mismatches with their new primer pair, and confirmed for at least one species. Other species not detected may lack sequenced representatives in GenBank with sufficiently long sequences, or even any sequenced representatives, which would have prevented taxonomic assignment. This could also be the case for very small dinoflagellates, and other very tiny, less-well studied taxa.

Overall, they concluded their full-length 18S rDNA metabarcoding approach works well for investigating marine protist communities, and that it can be successfully achieved using Nanopore sequencing.

A great step forward in marine plankton metabarcoding!
📄 Gaonkar, C. C., & Campbell, L. (2024). https://doi.org/10.1002/ece3.11232

📷 Diatoms through the microscope by Prof. Gordon T. Taylor, Stony Brook University

Here’s a free downloadable costing & budgeting spreadsheet tool to help you fully cost your PCR lab work; calculate what you need to buy (and how much); or to help you plan what to take on fieldwork.

Great for PCR beginners, and hopefully useful for the more experienced too! 👇

It follows a simple process, and probably a very obvious one to anyone who does a lot of budgeting:

⭐ Break down your PCR workflows into single steps, and put each step in a different spreadsheet row

⭐ Make a list of all the items used in that workflow along the top of the spreadsheet to form a table

⭐ Count each use of each item for each step, and enter those into the table

⭐ Sum the uses of each item (as a pipette tip or µL of a reagent) at the bottom

And you’ve now got a detailed count of all the items that you need, in the quantities that you need them.

You can then work out the costs by:

💲 Entering the price of each item and its volume or number of units
💲 Calculating the cost of each unit of each item
💲 Calculating the cost of all the units of each item used in the workflow.

You can use this information to calculate the cost of a single DNA extraction or PCR, a batch of PCRs, the amount of each item needed for a given number of samples, or the startup costs for a full set of reagents and consumables.

Our spreadsheet tool covers a range of examples using Bento Lab workflows, reagents, and consumables, but it's easy to delete all of this data and put in your own protocols, items, and prices to make it reflect your workflows

If you find this method and spreadsheet useful, please share it around to anyone else you think it may help!

📄 You can find it here: https://bento.bio/resources/bento-lab-advice/free-pcr-workflow-budgeting-spreadsheets/

Today is , commemorating the discovery of the structure of DNA in 1953 & completion of the Human Genome Project in April 2003.

It's also a good day to celebrate PCR, so here's an easy-reading history of the polymerase chain reaction plus a few landmark articles 👇

🧬The main article we’d like to share is a review of the development of PCR by Zhu et al. (2020), which is a straightforward but detailed read that might be particularly useful for students and PCR beginners:

Zhu et al. (2020). PCR past, present and future. Biotechniques, 69(4), 317-325.
https://future-science.com/doi/full/10.2144/btn-2020-0057

The article is also a great source for references for some important PCR breakthroughs, such as...

🧬The landmark paper in which PCR was first described, by Saiki et al. (1985):

Saiki et al. (1985). Enzymatic amplification of β-globin genomic sequences and restriction site analysis for diagnosis of sickle cell anemia. Science, 230(4732), 1350-1354.http://mun.ca/biology/scarr/Saiki,_Mullis_et_al_1985_Science_230,1250.pdf

🧬The first uses of a polymerase enzyme that could tolerate thermal cycling (Taq polymerase, from the thermophilic bacterium Thermus aquaticus):

Saiki et al. (1988). Primer-directed enzymatic amplification of DNA with a thermostable DNA polymerase. Science, 239(4839), 487-491.
https://citeseerx.ist.psu.edu/document?repid=rep1&type=pdf&doi=5fd581fcb0a18755701d94f99672eb568babdf82

🧬One of the first applications of multiplex PCR, by Chamberlain et al. (1988):

Chamberlain et al. (1988). Deletion screening of the Duchenne muscular dystrophy locus via multiplex DNA amplification. Nucleic acids research, 16(23), 11141-11156.https://academic.oup.com/nar/article-pdf/16/23/11141/3942426/16-23-11141.pdf

🧬The development of DNA fingerprinting from trace amounts of DNA, a major step forward in the use of PCR in forensics:

Jeffreys et al. (1988). Amplification of human minisatellites by the polymerase chain reaction: towards DNA fingerprinting of single cells. Nucleic Acids Research, 16(23), 10953-10971.https://academic.oup.com/nar/article-pdf/16/23/10953/3941961/16-23-10953.pdf

🧬And the use of PCR to detect infectious disease-causing agents in humans, such as HIV:

Kwok et al. (1987). Identification of human immunodeficiency virus sequences by using in vitro enzymatic amplification and oligomer cleavage detection. Journal of virology, 61(5), 1690-1694.
https://journals.asm.org/doi/pdf/10.1128/jvi.61.5.1690-1694.1987

It’s notable that all of these examples were published only two or three years after the invention of PCR. It must have seemed like a method with incredible potential at the time! (And it still is!)

Happy !

Photo: Raymond Gosling/King's College London - http://www-project.slac.stanford.edu/wis/images/photo_51.jpg

Here's a great direct PCR DNA barcoding approach for communities of cryptograms (e.g. of lichens, non-lichenised fungi, bryophytes, microalgae, biocrusts, or biofilms). It allows the simultaneous processing of 4 different taxonomic groups in ~1 hr 6 mins!

The method, by Jung et al. (2024), involves:

A direct PCR approach to allow fine-resolution sampling of tiny organisms, such as the sampling 0.2-2 mm pieces of individual colonies from a piece of desert grit crust stone

A direct PCR kit that includes lysis buffers, proteinase K, and a fast PCR-inhibition-resistant polymerase (Platinum Direct PCR Universal Master Mix Kit)

Using the same PCR thermocycler programs (but different primers) for all four organismal groups (lichens and non-lichenised fungi, cyanobacteria, green algae, and bryophytes)

Using this method, the authors report a high (>90%) success rate for successful amplification and high-quality Sanger sequencing, for various gene regions, from all organismal groups investigated.

Additionally, their estimated time from sampling preparation to PCR product was only 1 hr 6 minutes, compared to up to 8 hrs for other similar published methods doing the same kinds of research.

They found that the biggest advantage of this method is that it can be used with extremely small amounts of biomass, such as tiny parts of colonies picked with tweezers under a dissecting microscope.

This could make it much easier to DNA barcode many smaller species, and allow investigation of the identities and interactions of cryptograms on a much smaller spatial scale, for example within lichen symbioses or soil crusts.

Another big advantage of this method is that specimens of different organismal groups can be sampled and prepped for direct PCR at the same time, and DNA barcodes amplified in the same PCR run.

This could save a lot of time and effort and make it easier to sample larger numbers of species — potentially very useful when dealing with crowded cryptogram communities containing many species!

Their method could also be ideal for fieldwork involving cryptogram DNA barcoding or metabarcoding thanks to its simplicity and speed, and the fast multi-taxon PCR could save precious electricity or generator petrol if working off-grid.

However, the authors did have some problems when working with some lichen groups (presumably due to PCR-inhibiting pigments), so they recommend that their method should be tested and evaluated whenever new organisms or primers are used.

A very useful method! It might also be worth trying parts of this method (e.g. combined taxon thermocycling, or direct PCR) even with standard reagents, just to see if they can be made to work...

You can read more about their methods and results here:

Jung et al. (2024). A direct PCR approach with low-biomass insert opens new horizons for molecular sciences on cryptogam communities.
https://journals.asm.org/doi/pdf/10.1128/aem.00024-24

Photo by Annette Meyer.

Why are red onions red, and white onions white, and can you detect this difference using ?

You can find out the answer in this detailed lab practical for teachers and students.

One great thing about this practical lesson is that it’s much more than just DNA extraction and a single PCR assay.

Instead, it leads the students through a PCR-based investigation of the different enzymes in the red pigment biosynthesis pathway to find out which one is defective.

It also describes a range of additional experiments that could be done — which are great to know about in terms of learning about the potential of PCR, , plant molecular biology, and plant breeding, even if you don’t do these experiments in practice.

Not bad for a lesson based around an everyday vegetable from the grocery store!

Briju & Wyatt (2015). Grocery store genetics: A PCR-based genetics lab that links genotype to phenotype. The American Biology Teacher, 77(3), 211-214.

https://researchgate.net/publication/279287533_Grocery_Store_Genetics_A_PCR-Based_Genetics_Lab_that_Links_Genotype_to_Phenotype

For anyone wanting to learn or teach basic phylogenetics for DNA barcoding and identification, here are three introductory tutorials for identifying fungi, tardigrades, and flies, all using the free MEGA package for molecular evolutionary genetics analysis.

All three tutorials cover essentially the same processes but are slightly different, so you could compare, contrast, find the one that works best for you, or combine the most useful parts of each!

🍄 The first resource, by Casanova et al. (2021) is an article describing three 2.5 hr lab practicals involving a mystery fungal specimen.

It covers DNA extraction using a spin column kit, amplification of the internal transcribed spacer (ITS) barcode region for fungi, and sequencing. The mystery fungus sequence is then identified using BLAST searching in GenBank and MEGA is used to produce a maximum parsimony phylogenic tree. Supporting files can be found in the supplementary data.

If you want to try this yourself without a spin column kit, it may be useful to know our Dipstick DNA Extraction Kit works very well for fungi and only takes a minute or less for extraction.

Suárez Casanova, V. M., & Shumskaya, M. (2021). Exploring DNA in biochemistry lab courses: DNA barcoding and phylogenetic analysis. Biochemistry and Molecular Biology Education, 49(5), 789-799.
https://iubmb.onlinelibrary.wiley.com/doi/full/10.1002/bmb.21551

⭐The second guide is a detailed protocol for tardigrade barcoding, by D’Elia et al. 2023.

It describes how to find, sample, extract DNA using a spin column kit, and amplify the 18s rRNA barcode region using tardigrade-specific primers. It then uses the DNA Subway online portal (by the DNA Learning Center, Cold Spring Harbors) to produce consensus sequences; uses BLAST searching in GenBank for an initial identification and download reference sequences, and finally MEGA to construct a multiple sequence alignment and phylogenetic tree to analyse the sequences.

For those without access to spin column kits and tardigrade-specific primers, you can also extract tardigrade DNA using the HotSHOT DNA extraction method (reducing the volumes to 20 µL), and use universal COI primers for barcoding.

Additionally, if you want to work through the analysis part without doing the DNA extraction and PCR, you could use the sequence here as the mystery sample: https://ncbi.nlm.nih.gov/nuccore/MH664934.1

D'Elia et al. 2023. A Complete Guide to Tardigrade Isolation and Phylogenetic Characterization for Undergraduate Students. https://dx.doi.org/10.17504/protocols.io.81wgb64qqlpk/v1

🪰 The third guide, by Lorusso et al. (2022), is a remote exercise working only with phylogenetics, which aims to guide a learner in the forensic sciences in constructing phylogenetic trees to identify unknown specimens.

In this case the mystery specimen is a fly associated with a decomposing body (because different flies are associated with different decomposition stages), but exactly the same process can be used for biodiversity surveys.

This exercise has a walkthrough of searching a COI barcode sequence in the BOLD database for an initial identification, followed by constructing a multiple sequence alignment and a phylogeny in MEGA, after which the initial identification can be reassessed.

Most of the exercise information is in the downloadable teaching materials, consisting of a Powerpoint presentation, a student handout with instructions, and reference sequences.

Lorusso et al. (2022). Applying phylogenetic tree building in MEGA X to forensic applications for identifying unknown specimens. (Version 2.0). QUBES Educational Resources. doi:10.25334/XY5F-XQ54
https://qubeshub.org/publications/2567/2

If you find any of these learning resources useful, or if you can share other similar examples, please do let us know!

For anyone learning or teaching pipetting, here are four great exercises for accurate pipetting and dilution skills, including one with a glucose monitor (very cool!), a simple dye-based exercise, pipetting incorrectly on purpose, and a friendly classroom “Pipetting Olympics”!

📍 Burnette et al. (2016) describe a simple and colourful pipetting practice exercise for high school, undergraduates, and adults in continuing education.

This practical uses a variety of food colouring dyes, a micropipette, and 1.5 mL tubes. Various volumes of different colours are pipetted into a tube, and the accuracy of pipetting can be assessed by comparison with a reference tube.

Burnette et al. 2016. Dilution and Pipetting Lesson Using Food Dyes. CourseSource. https://qubeshub.org/community/groups/coursesource/publications?id=2556&v=1

📍 Mel et al. (2019) describe a pipetting exercise that demonstrates the difference in volumes when pipetting accurately and inaccurately.

Students are guided to weigh an empty 1.5 mL tube on an analytical balance, pipette 60 µL or 120 µL of water into it after pipetting using the first (correct) or second stop (incorrect) on a micropipette; weigh the full tubes; and then compare the two volumes and pipetting methods with a statistical analysis (a T-test, although ANOVA could also be used).

Mel, S.F., Micou, M.K., Gaur, K., Lenh, D., Liu, C.Z., and Lo, S.M. 2019. Learning to Pipet Correctly by Pipetting Incorrectly? CourseSource. https://qubeshub.org/community/groups/coursesource/publications?id=2620&v=1

📍 Jawad et al. (2021) describe a very cool remote learning exercise (developed during the COVID-19 pandemic) that can be done at home using cheap glucose monitors and glucose/bovine serum albumin solutions.

In this exercise, students investigate their pipetting accuracy using a small milligram scale, and then create a dilution calibration curve using a glucose monitor. They also investigate the concepts of standard deviation and standard error and apply these to their data.

This exercise also has a comprehensive handout in the supplementary data, including a complete walkthrough and also questions for the student to answer.

Jawad et al. (2021). Remote laboratory exercise to develop micropipetting skills. Journal of Microbiology & Biology Education, 22(1), 10-1128. https://www.sciencedirect.com/org/science/article/pii/S1935787721000769

📍 Richter et al. (2022) describe an exercise they call the “Pipetting Olympics”, in which students can compete (in a friendly manner of course!) to accurately fill pipette various dilutions of a dye into a 96-well plate, and then have their results assessed using a plate reader.

Richter et al. (2022). Pipette Olympics: An Engaging Exercise for Undergraduate Laboratory Training. Journal of Undergraduate Neuroscience Education, 21(1), A81. https://www.ncbi.nlm.nih.gov/pmc/articles/PMC10558238/

If you find any of these useful, or have other pipetting exercises you would recommend, please let us know!

DNA sampling by drone?

Team Limelight Rainforest are using aerial robots from Outreach Robotics (and a Bento Lab!) to sample and DNA barcode insects in the Amazon basin in Ecuador.

They're on Kickstarter NOW aiming to catalog 5000 insect taxa. Support them here: https://www.kickstarter.com/projects/599390557/limelight-rainforest-5k

Sampling using aerial drones is a clever approach to less invasive monitoring that project leader Dr Thomas Walla at Colorado Mesa University believes will lead to more rapid collection of biodiversity data: "This is a project with the intention of dramatically increasing the rate of biodiversity surveying in the Amazon rainforest".

Limelight Rainforest 5K - Race to Identify Race with us to fund 5000 insect identifications in the Amazon!