A scientific article led by my colleague Dr. Alaa Rabee at the Desert Research Center in Egypt was just published online and is now available! Dr. Rabee and I have been collaborating remotely on projects related to the bacteria in the rumen of camels, sheep, and cows, as Dr. Rabee’s work focuses on the isolation of bacteria which can degrade plant materials efficiently and could be used to produce biofuels. He will be spending 6 months working in my lab as a visiting scholar, which was delayed until this year because of the pandemic.
Rabee, A.E., Sayed Alahl, A.A., Lamara, M., Ishaq, S.L. 2022. Fibrolytic rumen bacteria of camel and sheep and their applications in the bioconversion of barley straw to soluble sugars for biofuel production. PLoS ONE 17(1): e0262304. Article.
Lignocellulosic biomass such as barley straw is a renewable and sustainable alternative to traditional feeds and could be used as bioenergy sources; however, low hydrolysis rate reduces the fermentation efficiency. Understanding the degradation and colonization of barley straw by rumen bacteria is the key step to improve the utilization of barley straw in animal feeding or biofuel production. This study evaluated the hydrolysis of barley straw as a result of the inoculation by rumen fluid of camel and sheep. Ground barley straw was incubated anaerobically with rumen inocula from three fistulated camels (FC) and three fistulated sheep (FR) for a period of 72 h. The source of rumen inoculum did not affect the disappearance of dry matter (DMD), neutral detergent fiber (NDFD). Group FR showed higher production of glucose, xylose, and gas; while higher ethanol production was associated with cellulosic hydrolysates obtained from FC group. The diversity and structure of bacterial communities attached to barley straw was investigated by Illumina Mi-Seq sequencing of V4-V5 region of 16S rRNA genes. The bacterial community was dominated by phylum Firmicutes and Bacteroidetes. The dominant genera were RC9_gut_group, Ruminococcus, Saccharofermentans, Butyrivibrio, Succiniclasticum, Selenomonas, and Streptococcus, indicating the important role of these genera in lignocellulose fermentation in the rumen. Group FR showed higher RC9_gut_group and group FC revealed higher Ruminococcus, Saccharofermentans, and Butyrivibrio. Higher enzymes activities (cellulase and xylanase) were associated with group FC. Thus, bacterial communities in camel and sheep have a great potential to improve the utilization lignocellulosic material in animal feeding and the production of biofuel and enzymes.
A few weeks ago, I sat down with Sheba A-J, one of the producers of the WeTalkScience podcast, to talk about one of my recent publications in the research journal iScience, at which Sheba is also an editor. Listen to find out how lobsters are like humans, how I got involved on a project working with ants and nematodes, and how you can help make science a more welcoming place.
The full publication is:
Ishaq, S.L., A. Hotopp, S. Silverbrand, J.E. Dumont, A. Michaud, J. MacRae, S. P. Stock, E. Groden. 2021. Bacterial transfer from Pristionchus entomophagus nematodes to the invasive ant Myrmica rubra and the potential for colony mortality in coastal Maine. iScience 24(6):102663. Article.
A collaborative paper on bacterial transfer in insects and the possible ecological impacts of that in the wild has been published in iScience! This work began a decade ago in the labs of Dr. Ellie Groden, recently retired Professor of Entomology in the School of Biology and Ecology at the University of Maine, and later Dr. Patricia Stock, a Professor in the School of Animal and Comparative Biomedical Sciences at the University of Arizona, who were investigating colony collapse of European fire ants (Myrmica rubra) which are invasive to Maine. The ants have a nasty bite, and can dramatically disturb the local plant and insect wildlife in coastal Maine.
When these invasive ant colonies collapsed, Drs. Groden and Stock wanted to find out why, as a possible means of developing a biological control strategy. It was thought that particular nematodes would ingest soil bacteria, and transfer it to ants once the worms invaded ant tissues to complete parts of their life cycle. This particular worm infection doesn’t kill the ants, but perhaps the soil bacteria were. Ants were collected from different colony sites, and investigations on the nematode worms inhabiting the ants were conducted.
Most of the work for this project was completed several years ago, with the exception of DNA sequencing data from a bacterial transfer experiment. I was added to the project by my collaborator at UMaine, Dr. Jean MacRae, an Associate Professor in the Department of Civil and Environmental Engineering who introduced me to the research team and shared the 16S rRNA dataset to use in my AVS 590 data analysis class in spring 2020. That semester was when the pandemic hit, and forced the course to move to remote-only instruction in March. UMaine graduate students Alice Hotopp and Sam Silverbrand were taking the class and learning 16S analysis on this dataset, and I mentored them through the analysis all the way to manuscript writing despite the incredible challenges that spring threw our way.
At the completion of the course, we shared the draft manuscript with the rest of the research team, who mentioned that several undergraduate honor’s theses had been written about the earlier experiment, but never published in a scientific journal. The team spent summer 2020 combining the three papers into one massive draft. The pandemic slowed down manuscript review, understandably, but I’m pleased to say that it was accepted for publication! In addition, this collaboration has led to further collaborations in the Ishaq Lab, several presentations (listed below), and is Sam’s first scientific publication, congrats Sam!!
Alice Hotopp, A., Samantha Silverbrand, Suzanne L. Ishaq, Jean MacRae, S. Patricia Stock, Eleanor Groden. “Can a necromenic nematode serve as a biological Trojan horse for an invasive ant?” Ecological Society of America 2021 (virtual). Aug 2-6, 2021 (accepted poster).
Ishaq*, S.L., Hotopp, A., Silverbrand, S., MacRae, J., Stock, S.P., Groden, E. “Can a necromenic nematode serve as a biological Trojan horse for an invasive ant?” Entomological Society of America 2020 (virtual). Nov 15-25, 2020. (invited talk)
The necromenic nematode Pristionchus entomophagus has been frequently found in nests of the invasive European ant Myrmica rubra in coastal Maine, United States, and may contribute to ant mortality and collapse of colonies by transferring environmental bacteria. Paenibacillus and several other bacterial species were found in the digestive tracts of nematodes harvested from collapsed ant colonies. Serratia marcescens, Serratia nematodiphila, and Pseudomonas fluorescens were collected from the hemolymph of nematode-infected wax moth (Galleria mellonella) larvae.
Virulence against waxworms varied by site of origin of the nematodes. In adult nematodes, bacteria were highly concentrated in the digestive tract with none observed on the cuticle. In contrast juveniles had more on the cuticle than in the digestive tract. . Host species was the primary factor affecting bacterial community profiles, but Spiroplasma sp. and Serratia marcescens sequences were shared across ants, nematodes, and nematode-exposed G. mellonella larvae.
In October 2017, Dr. Rich Corsi came to visit Oregon for two weeks during a sabbatical from the University of Texas, Austin. During his stay, Rich and I, and other BioBE/ESBL researchers chatted about doing a pilot study that would bring UT’s indoor chemistry work together with BioBE’s indoor microbial work.
Since we began our collaboration in the fall of 2017, only one of the research team is still in their original position (Jeff Kline at ESBL)! Rich Corsi, Ying Xu, and myself have all gone on to faculty positions elsewhere, graduate student Chenyang Bi defended and started a post-doc position, and the two undergrads working with me, Susie Nunez and Samantha Velazquez, graduated and went on to other things! Science collaborations work best when they can stand the test of time and geography. The benefit to everyone moving around is that you are able to hold collaboration meetings in new and exciting places each time.
In addition to the literature review we collaborated on, we eventually did get a research pilot running, and it has now been published in PeerJ!
Chenyang Bi 2,3, Jeff Kline 1,4, Susie Nunez1,
Richard Corsi 3,5, Ying Xu 3,6, Suzanne L. Ishaq1,7*
1 Biology and
the Built Environment Center, University
of Oregon, Eugene, OR, 97403
of Civil Environmental Engineering, Virginia Polytechnic Institute and State University,
Blacksburg, VA 24061 (current)
of Civil, Architectural and Environmental Engineering, University of Texas,
Austin, TX 78712
Studies and Buildings Laboratory, University of Oregon, Eugene, OR, 97403
Maseeh College of Engineering and Computer Science, Portland State University,
Portland, OR 97207 (current)
6 Department of Building Science, Tsinghua University,
100084, Beijing, P. R. China (current)
7 School of Food and Agriculture, University of Maine, Orono, ME 04469 (current)
phthalate (DEHP) is a plasticizer used in consumer products and building
materials, including polyvinyl chloride flooring material. DEHP adsorbs from
material and leaches into soil, water, or dust, and presents an exposure risk
to building occupants by inhalation, ingestion, or absorption. A number of bacterial isolates are
demonstrated to degrade DEHP in culture, but bacteria may be susceptible to it
as well, thus this study examined the relation of DEHP to bacterial communities
in dust. Polyvinyl chloride flooring was
seeded with homogenized house dust and incubated for up to 14 days, and
bacterial communities in dust were identified at days 1, 7, and 14 using the
V3-V4 regions of the bacterial 16S rRNA gene.
DEHP concentration in dust increased over time, as expected, and
bacterial richness and Shannon diversity were negatively correlated with DEHP
sequence variants of Bacillus, Corynebacterium jeddahense, Streptococcus, and Peptoniphilus were relatively more abundant at low concentrations
of DEHP, while some Sphingomonas, Chryseobacterium, and a member of the
Enterobacteriaceae family were relatively more abundant at higher
concentrations. The built environment is
known to host lower microbial diversity and biomass than natural environments,
and DEHP or other chemicals indoors may contribute to this paucity.
After several years of bouncing through internal and external review, I’m pleased to announce that the first microbes paper out of the Montana State University Fort Ellis project has been published in Geoderma! The Fort Ellis research has encompassed multiple labs, projects, and many personnel, as it was a large collaboration looking at the effect of different farming systems on biodiversity at the macro (plant), mini (insect), and micro (-be) levels. Spanning multiple years, this project has been a massive undertaking that I briefly participated in but anticipate getting four publications out of (two more are in preparation).
Despite knowledge that management practices, seasonality, and plant phenology impact soil microbiota; farming system effects on soil microbiota are not often evaluated across the growing season. We assessed the bacterial diversity in soil around wheat roots through the spring and summer of 2016 in winter wheat (Triticum aestivium L.) in Montana, USA, from three contrasting farming systems: a chemically-managed no-tillage system, and two USDA-certified organic systems in their fourth year, one including tillage and one where sheep grazing partially offsets tillage frequency. Bacterial richness (range 605 – 1174 OTUs) and evenness (range 0.80 – 0.92) peaked in early June and dropped by late July (range 92 – 1190, 0.62-0.92, respectively), but was not different by farming systems. Organic tilled plots contained more putative nitrogen-fixing bacterial genera than the other two systems. Bacterial community similarities were significantly altered by sampling date, minimum and maximum temperature at sampling, bacterial abundance at date of sampling, total weed richness, and coverage of Taraxacum officinale, Lamium ampleuxicaule, and Thlaspi arvense. This study highlights that weed diversity, season, and farming management system all influence soil microbial communities. Local environmental conditions will strongly condition any practical applications aimed at improving soil diversity, especially in semi-arid regions where abiotic stress and seasonal variability in temperature and water availability drive primary production. Thus, it is critical to incorporate or address seasonality in soil sampling for microbial diversity.
I’m pleased to announce that the “particle size” project is officially published! I inherited this dataset of bacterial 16S rRNA sequences in 2015, while working for the Yeoman Lab. This collaborative project combined nutrition, animal production, and microbial ecology to look at the effect of diet particle size on lambs and their rumen bacteria. While small in size, the project was large in scope – despite everything we know about how different diet components encourage different microbial communities to survive in the digestive tract, we know practically nothing about how the size of the particles in that diet might contribute.
A lot of factors can be manipulated to help get the most out of one’s diet, including adjusting ingredients for water content, palatability, ease of chewing, and how easy the ingredients are to digest. For example, highly fibrous foods with larger particles/pieces require more chewing, as well as a longer time spent in the rumen digesting so that microorganisms have plenty of time to break the chemical bonds of large molecules. Smaller food particles can reduce the time and effort spent chewing, allow for more surface area on plant fibers for microorganisms to attach to and digest faster, and speed up the movement of food through the digestive tract. On the other hand, moving food too quickly could reduce the amount of time microorganisms can spend digesting, or time the ruminant can absorb nutrients across their GI tract lumen, or cause slow-growing microbial species to wash out.
Suzanne L. Ishaq1, Medora M. Lachman2, Benjamin A. Wenner3, Amy Baeza2, Molly Butler2, Emily Gates2, Sarah Olivo1, Julie Buono Geddes2, Patrick Hatfield2, Carl J. Yeoman2
Biology and the Built Environment Center, University of Oregon, Eugene, Oregon, United States of America
Department of Animal and Range Sciences, Montana State University, Bozeman, Montana, United States of America
Department of Animal Sciences, The Ohio State University, Columbus, Ohio, United States of America
Diet composed of smaller particles can improve feed intake, digestibility, and animal growth or health, but in ruminant species can reduce rumination and buffering – the loss of which may inhibit fermentation and digestibility. However, the explicit effect of particle size on the rumen microbiota remains untested, despite their crucial role in digestion. We evaluated the effects of reduced particle size on rumen microbiota by feeding long-stem (loose) alfalfa hay compared to a ground and pelleted version of the same alfalfa in yearling sheep wethers during a two-week experimental period. In situ digestibility of the pelleted diet was greater at 48 h compared with loose hay; however, distribution of residual fecal particle sizes in sheep did not differ between the dietary treatments at any time point (day 7 or 14). Both average daily gain and feed efficiency were greater for the wethers consuming the pelleted diet. Observed bacterial richness was very low at the end of the adaptation period and increased over the course of the study, suggesting the rumen bacterial community was still in flux after two weeks of adaptation. The pelleted-hay diet group had a greater increase in bacterial richness, including common fibrolytic rumen inhabitants. The pelleted diet was positively associated with several Succiniclasticum, a Prevotella, and uncultured taxa in the Ruminococcaceae and Rickenellaceae families and Bacteroidales order. Pelleting an alfalfa hay diet for sheep does shift the rumen microbiome, though the interplay of diet particle size, retention and gastrointestinal transit time, microbial fermentative and hydrolytic activity, and host growth or health is still largely unexplored.
Last year, one of my former research groups at Montana State University was awarded a USDA NIFA Foundational program grant, and I am a sub-award PI on that grant. We’ll be working together to investigate the effect of diversified farming systems – such as those that use cover crops, rotations, or integrate livestock grazing into field management – on crop production and soil bacterial communities: “Diversifying cropping systems through cover crops and targeted grazing: impacts on plant-microbe-insect interactions, yield and economic returns.”
The first soil samples were collected in Montana this summer, and I have been processing them for the past few weeks. I am using the opportunity to train a master’s student on microbiology and molecular genetics lab work.
Tindall Ouverson started this fall as a master’s student at MSU, working with Fabian Menalled and Tim Seipel in Bozeman, MT. She’s an environmental and soil scientist, and this is her first time working with microbes. She was here in Eugene for just a few days to learn everything needed for sequencing: DNA extraction, polymerase chain reaction, gel electrophoresis and visualization, DNA cleanup using magnetic beads, quantification, and pooling. Despite not having experience in microbiology or molecular biology, Tindall showed a real aptitude and picked up the techniques faster than I expected!
Once the sequences are generated, I’ll be (remotely) training Tindall on DNA sequence analysis. I’ll also be serving as one of her thesis committee members! Tindall will be the first of (hopefully) many cross-trained graduate students between myself and collaborators at MSU.
Zinc is an important mineral in your diet; it’s required by many of your enzymes and having too much or too little can cause health problems. We know quite a bit about how important zinc is to sheep, in particular for their growth, immune system, and fertility. We also know that organically- versus inorganically-sourced zinc differs in its bio-availability, or how easy it is for cells to access and use it. Surprisingly, we know nothing about how different zinc formulations might affect gut microbiota, despite the knowledge that microorganisms may also need zinc.
This collaborative study was led by Dr. Whit Stewart and his then-graduate student, Chad Page, while they were at Montana State University (they are now both at the University of Wyoming). Chad’s work focused on how different sources of zinc affected sheep growth and performance (previously presented, publication forthcoming), and I put together this companion paper examining the effects on rumen bacteria.
The pre-print is available now for Journal of Animal Science members, and the finished proof should be available soon. JAS is the main publication for the American Society of Animal Science, and one of the flagship journals in the field.
Zinc amino acid supplementation alters yearling ram rumen bacterial communities but zinc sulfate supplementation does not.
Ishaq, S.L., Page, C.M., Yeoman, C.J., Murphy, T.W., Van Emon, M.L., Stewart, W.C. 2018. Journal of Animal Science. Accepted.Article.
Despite the body of research into Zn for human and animal health and productivity, very little work has been done to discern whether this benefit is exerted solely on the host organism, or whether there is some effect of dietary Zn upon the gastrointestinal microbiota, particularly in ruminants. We hypothesized that 1) supplementation with Zn would alter the rumen bacterial community in yearling rams, but that 2) supplementation with either inorganically-sourced ZnSO4, or a chelated Zn amino acid complex, which was more bioavailable, would affect the rumen bacterial community differently. Sixteen purebred Targhee yearling rams were utilized in an 84 d completely-randomized design, and allocated to one of three pelleted dietary treatments: control diet without fortified Zn (~1 x NRC), a diet fortified with a Zn amino acid complex (~2 x NRC), and a diet fortified with ZnSO4 (~2 x NRC). Rumen bacterial community was assessed using Illumina MiSeq of the V4-V6 region of the 16S rRNA gene. One hundred and eleven OTUs were found with > 1% abundance across all samples. The genera Prevotella, Solobacterium, Ruminococcus, Butyrivibrio, Olsenella, Atopobium, and the candidate genus Saccharimonas were abundant in all samples. Total rumen bacterial evenness and diversity in rams were reduced by supplementation with a Zn-amino-acid complex, but not in rams supplemented with an equal concentration of ZnSO4, likely due to differences in bioavailability between organic and inorganically-sourced supplement formulations. A number of bacterial genera were altered by Zn supplementation, but only the phylum Tenericutes was significantly reduced by ZnSO4 supplementation, suggesting that either Zn supplementation formulation could be utilized without causing a high-level shift in the rumen bacterial community which could have negative consequences for digestion and animal health.
Sequence data contamination from biological or digital sources can obscure true results and falsely raise one’s hopes. Contamination is a persist issue in microbial ecology, and each experiment faces unique challenges from a myriad of sources, which I have previously discussed. In microbiology, those microscopic stowaways and spurious sequencing errors can be difficult to identify as non-sample contaminants, and collectively they can create large-scale changes to what you think a microbial community looks like.
Samples from large studies are often processed in batches based on how many samples can be processed by certain laboratory equipment, and if these span multiple bottles of reagents, or water-filtration systems, each batch might end up with a unique contamination profile. If your samples are not randomized between batches, and each batch ends up representing a specific time point or a treatment from your experiment, these batch effects can be mistaken for a treatment effect (a.k.a. a false positive).
“The times were statistically greater than prior time periods, while simultaneously being statistically lesser to prior times, according to longitudinal analysis.”
Over the past year, I analyzed a particularly complex bacterial 16S rRNA gene sequence data set, comprising nearly 600 home dust samples, and about 90 controls. Samples were collected from three climate regions in Oregon, over a span of one year, in which homes were sampled before and approximately six weeks after a home-specific weatherization improvement (treatment homes) or simply six weeks later in (comparison) homes which were eligible for weatherization but did not receive it. As these samples were collected over a span of a year, they were extracted with two different sequencing kits and multiple DNA extraction batches, although all within a short time after collection. The extracted DNA was spread across two sequence runs to allow for data processing to begin on cohort 1, while we waited for cohort 2 homes to be weatherized. Thus, there were a lot of opportunities to introduce technical error or biological contamination that could be conflated with treatment effects.
On top of this, each home was unique, with it’s own human and animal occupants, architectural and interior design, plants, compost, and quirks, and we didn’t ask homeowners to modify their behavior in any way. This was important, as it meant each of the homes – and their microbiomes – are somewhat unique. Therefore I didn’t want to remove sequences which might be contaminants on the basis of low abundance and risk removing microbial community members which were specific to that home. After the typical quality assurance steps to curate and process the data, which can be found on GitHub as an R script of a DADA2 package workflow, I needed to decide what to do with the negative controls.
Because sequencing is expensive, most of the time there is only one negative control included in sequencing library preparation, if that. The negative control is a blank sample – just water, or an unused swab – which does not intentionally contain cells or nucleic acids. Thus anything you find there will have come from contamination. The negative control can be used to normalize the relative abundance numbers – if you find 1,000 sequences in the negative control, which is supposed to have no DNA in it, then you might only continue looking at samples with a certain amount higher than 1,000 sequences. This risks throwing out valid sequences that happen to be rare. Alternatively, you can try to identify the contaminants and remove whole taxa from your data set, risking the complete removal of valid taxa.
I had three types of negative controls: sterile DNA swabs which were processed to check for biological contamination in collection materials, kit controls where a blank extraction was run for each batch of extractions to test for biological contamination in extraction reagents, and PCR negative controls to check for DNA contamination of PCR reagents. In total, 90 control samples were sequenced, giving me unprecedented resolution to deal with contamination. Looking at the total number of sequences before and after my quality-analysis processing, I can see that the number of sequences in my negative controls reduces dramatically; they were low-quality in some way and might be sequencing artifacts. But, an unsatisfactory number remain after QA filtering; these are high-quality and likely come from microbial contamination.
I wasn’t sure how I wanted to deal with each type of control. I came up with three approaches, and then looked at unweighted, non-rarefied ordination plots (PCoA) to watch how my axes changed based on important components (factors). What follows is a narrative summarize of what I did, but I included the R script of my phyloseq package workflow and workaround on GitHub.
“In microbial ecology, preprints are posted on late November nights. The foreboding atmosphere of conflated factors makes everyone uneasy.”
Ordination plots visualize lots of complex communities together. In both ordination figures below, each point on the graph represents a dust sample from one house. They are clustered by community distance: those closer together on the plot have a more similar community than points which are further away from each other. The points are shaped by the location of the samples, including Bend, Eugene, Portland, along with a few pilot samples labeled “Out”, and negative controls which have no location (not pictured but listed as NA). The points are colored by DNA extraction b
In Figure 1, the primary axis (axis 1) shows a clear clustering of samples by DNA extraction batch, but this is also mixed with geographic location, and as it turns out – date of collection and sequencing run. We know from other studies that geographic location, date of collection, and sequencing batch can all affect the microbial community.
Approach 1: Subtraction + outright removal
This approach subsets my data into DNA extraction batches, and then uses the number of sequences found in the negative controls to subtract out sequences from my dust samples. This assumes that if a particular sequence showed up 10 times in my negative control, but 50 times in my dust samples, that only 40 of those in my dust sample were real. For each of my DNA extraction batch negative control samples, I obtained the sum of each potential contaminant that I found there, and then subtracted those sums from the same sequence columns in my dust samples.
Approach 1 was alright, but there was still an effect of DNA extraction batch (indicated by color scale) that was stronger than location or treatment (not included on this graph). This approach is also more pertinent for working with OTUs, or situations where you wouldn’t want to remove the whole OTU, just subtract out a certain number sequences from specific columns. There is currently no way to do that just from phyloseq, so I made a work-around (see the GitHub page). However, using DADA2 gives you Sequence Variants, which are more precise and I found it’s better to remove them with approach 3.
Approach 2: Total Removal
This approach removes any contaminant sequences that is found in ANY of the negative controls from ALL the house samples, regardless of which negative control was for which extraction batch. This approach assumes that if it a sequence was found as a contaminant in a negative control somewhere, that it is a contaminant everywhere.
Once again, approach 2 was alright, and now that primary axis (axis 1) of potential batch effect is now my secondary axis; so there is still an effect of DNA extraction batch (indicated by color scale) but it is weaker. When I recolor by different variables, there is much more clustering by Treatment than by any batch effects. However, that second axis is also one of my time variables, so don’t want to get rid of all of the variation on that axis. But, since my negative kit controls showed a lot of variation in number and types of taxa, I don’t want to remove everything found there from all samples indiscriminately.
Additionally, I don’t favor throwing sequences out just because they were a contaminant somewhere, particularly for dust samples. Contamination can be situational, particularly if a microbe is found in the local air or water supply and would be legitimately found in house dust but would have also accidentally gotten into the extraction process.
Approach 3: “To each its own”
This approach removes all the sequences from PCR and swab contaminant SVs fully from each cohort, respectively, and removes extraction kit contaminants fully from each DNA extraction batch, respectively. I took all the sequences of the SVs found in my dust samples and made them into a vector (list), and then I took all the sequences of the SVs found in my controls and made them into a different vector. I effectively subtracted out the contaminant SVs by name, but asking to find the sequences which were different between my two lists (thus returning the sequences which were in my dust samples but not in my control samples). I did this respective to each sequencing cohort and batch, so that I only remove the pertinent sequences (ex. using kit control 1 to subtract from DNA extraction batch 1).
In Figure 4, potential batch effect is solidly my secondary axis and not the primary driving force behind clustering. The primary axis (axis 1) shows a clear separation by climate zone, or location of homes, once the batch contamination has been removed. When I recolor by different variables, there is much more clustering by Treatment and almost none by batch effects. I say almost none, because some of my DNA extraction batches also happen to be Treatment batches, as they represent a subset of samples from a different location. Thus, I can’t tell if those samples cluster separately solely because of location or also because of batch effect. However, I am satisfied with the results and ready to move on.
Unlike its namesake, this tale has a happier ending.
Last night I participated in the Oregon Museum of Science and Industry (OMSI) After Dark event: “It’s Alive! (Mind and Body)”. OMSI regularly puts on After Dark events, where adults can check out the museum, listen to lectures in the planetarium, and engage in interactive science experiments and activities, all while enjoying an open bar. Last night, I had a great time giving a short presentation on “Ishaq OMSI After Dark 20180425“!