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Vampire Ventures

Oct 30, 2014
Michelle Tetreault Carlson

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Two vampires walk into a bar and call for the bartender.

“I’ll have a glass of blood,” says one.

“I’ll have a glass of plasma,” says the other.

“Okay,” replies the bartender, “That’ll be one blood and one blood lite.”

(insert laughter here)

 

Yep, we are celebrating the scary season and launching our new PowerMag Blood DNA/RNA Isolation kit!

Blood:  What’s in it?

Blood contains a mixture of plasma, red and white cells and platelets.  It is a unique beast among sample types, because while the quantity of nucleic acids in blood is copious, this genetic gold mine bathes inside a complex soup of cellular debris and protein. These contaminants can interfere with downstream PCR and sequencing.  It is the hemoglobin in particular, within the red cells, which causes major issues in DNA/RNA contamination and PCR inhibition.

Fortunately, dirty samples don’t scare us at here at MO BIO.  After all, we know how to get clean nucleic acids from feces and soil, no problemo!

Blood Collection and Storage:

Blood should be collected into an anticoagulant coated tube.  Otherwise, the blood will clot and bind up most of your DNA/RNA containing cells.  Standard blood collection tubes typically contain EDTA or citrate to prevent this.  Neither of these interferes with downstream genetic analysis.  Heparin can also prevent clotting but because it tends to bind to DNA, we don’t recommend it.

If whole blood can’t be processed right away it’s okay to store it at 4oC for up to two weeks for DNA isolation or for an hour or two for RNA isolation.  Any longer than this and you’ll need to take alternative action.   If you only want DNA you can simply freeze the blood in small aliquots (250-500 μL) at -20oC or -80oC for long term storage.

For RNA extraction from blood, you’ll need to be more vigilant.  RNases are tough enzymes and they can continue to have activity even while frozen.  If the blood cannot be processed quickly then we recommend collecting the blood in an RNA stabilization buffer such as that contained in PAXgene™ Blood RNA Tubes.  In these tubes, intracellular RNA will be stable for three days at 18 to 25°C or five days at 2 to 8°C.

Another option, if you can’t get to your RNA prep right away is to isolate, lyse and freeze the white cell pellet in an RNase inhibiting buffer.   For example, the white pellet can be stored in the PowerMag® WBC Lysis solution contained in the PowerMag Blood DNA/RNA Isolation kit. (See below)  Store the pellets at -20oC or -80oC. Once you are ready to extract, bring the sample to room temperature and proceed with the rest of the protocol.

Getting DNA & RNA from Blood13814735_s

In our PowerMag Blood DNA/RNA Isolation kit, the lysis of RBC and WBC is buffer based and makes use of the fact that in mammalian blood, genomic DNA (gDNA) is only contained in the white cells (leukocytes).  The Red cells (erythrocytes) and platelets lack a nucleus and so neither contains gDNA.  (bird erythrocytes are an exception)   In general, the average number of white cells in 1 mL of human blood is about 7 million.

The first step in our protocol uses a hypotonic lysis buffer that preferentially lyses RBC.  The WBC are pelleted and then the heme containing RBC supernatant is removed.  A chaotrophic buffer is added to the WBC pellet and the nucleic acids are released.  As mentioned above, this is a good point to freeze the sample for future RNA isolation at (-20 C or -80 C) for long term storage if the sample can’t be processed right away.  Our ClearMag® Beads are then used to capture the nucleic acids without binding unwanted contaminants.   Subsequent washes and elution generate ready-to-use DNA and RNA for most any downstream application.

Average RNA and DNA yields?

Not all blood samples are the same. They differ in the number of WBCs and this affects the quantity of nucleic acids isolated from each sample. WBC count can vary based on the health of the subject at the time of blood sampling. Nucleic acid yields vary by species as well.  However, average yields for MO BIO employees were 1 microgram of RNA and 3 micrograms of DNA from 200 microliters of whole (non-vampire) blood.

High Molecular Weight nucleic acids

If you are looking for high molecular weight DNA from blood in a high throughput manner, this kit is the way to go!  We’ve been able to isolate up to 100kb fragments of DNA so if you are looking for targets that are low in abundance or need longer chunks of DNA for sequencing purposes, look to this blood kit!

With all of that said and the vampire in R&D, hunger is looming and its best to get back to the bench where my stock of holy water and garlic bulbs are awaiting my arrival.  Later pumpkin skaters!

References:

  1. Yokota M, Tatsumi N, Nathalang O, Yamada T, Tsuda I. (1999). “Effects of Heparin on Polymerase Chain Reaction for Blood White Cells”. J. Clin. Lab. Anal. 13: 133–140.
  2. DNA isolation by a rapid method from human blood samples: effects of MgCl2, EDTA, storage time, and temperature on DNA yield and quality. Lahiri DK1, Schnabel B. Biochem Genet. 1993 Aug;31(7-8):321-8.

 

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Monster Cryo Tour

Core prep

 

Warning —> If you don’t use twitter, this article might give you the extra boost to go ahead start tweeting away..

I met Arwyn’s tweet one day and rest is history!  He had posted results from his undergraduate class about how there are essential oils such as lavendar that are actually a more effective bacteriocide than other toxic chemicals such as bleach.  Super Rad result!  I got in touch with him and, over time, learned about his incredible and very global climate change relevent research on microbes that live on ice and snow of the Arctic.

I sent him a few basic questions and voilà!  Enjoy.

1. Tell MO BIO about your project

My principal interests are in the interactions between microbes and glacial systems. This summer is very busy for my team as we are working on projects in Svalbard, the Canadian Arctic, Greenland, the Swedish Arctic and the European Alps. At the moment we are wrapping up our Greenland project, which is funded by the Royal Society in the UK. The project aims to build our understanding of how microbial communities on the surface of the Greenland ice sheet change through space and time. Here I have been working with my co-investigators Dr Tristram Irvine-Fynn, also of Aberystwyth University, and Dr. Joseph Cook of Derby University to collect samples and conduct experiments from a science camp on the ice sheet run by the Dark Snow Project.

Midnight Sampling

2. Why is it important?

Not too long ago scientists assumed that glaciers and ice sheets were too hostile for life to survive on them. We know now that microbial life is abundant, active and diverse in form upon, within and beneath glaciers and ice sheets. The microbial population of glacial systems is vast and poorly explored, but microbes play key roles in how glaciers and ice sheets work (DOI: 10.1002/wat2.1029). We recently published some papers on the abundance of microbes at the ice surface (e.g. DOI: 10.1002/cyto.a.22411). To put the numbers into context – well, the media seem to measure glacial disaster in SI units of “Manhattans”. I calculated that an iceberg the size of Manhattan would harbour an equivalent number of microbes in its top few metres of ice as the number of cells in the human population of Manhattan. Just a decade or so ago we had little idea these were habitats for life.
The flipside of this is that glacier melting becomes an ecological problem and not “just” a case of rising sea levels. Microbial habitats on the ice surface are both vulnerable to climate change and amplify its effects. Microbial biofilms and aggregates at the glacier surface, for example cryoconite and ice or snow algae, darken the ice, reducing its albedo and promote melt. In the bluntest of terms the microbes represent a powerful biological feedback to melting at the surface of glaciers and the Greenland ice sheet. We can see “dark zones” of low albedo and intense melting from satellite observations. When you ground-truth these areas you find that these areas are rich in cryoconite ecosystems, and that the “darkness” is derived from humic compounds made by microbes in the cryoconite.

3. What are some potential outcomes?

I hope our project will help map how these microbial communities change over space and time, in particular in relation to melting seasons. Although as I mentioned earlier, we are focused on how microbes cause melting, microbes also respond to melting, and that’s an important facet of our work. Back in 2012, 97% of the surface area of the Greenland ice sheet experienced surface melting, if only for a few days or a week at its extreme limits. On the basis of work on Svalbard glaciers, I hypothesized (doi:10.1038/ismej.2013.51) that some bacterial populations could respond to this melting episode, creating a spatially-expansive but very brief microbial bloom across Greenland in response to the availability of liquid water and nutrients. A massive event in nature but because of its invisible microbial nature, two years later we have no idea if it really happened or not. Last Saturday we lucked out and were able to take samples far inland to see if there are any traces of microbial changes in response to previous melt episodes. I can’t wait to get these samples back to my lab in Aberystwyth.

4. How is your science important to the public?

My team is intensely passionate about how microbes interact with the ice surface. What began as an academic pursuit for us has assumed a broader significance because of the feedbacks between biology and ice melting. We know as the climate warms, Earth’s glaciers and ice sheets will contribute to raising sea levels. This will affect humans across the world, be it people living in low-lying coastal areas which will be at risk of inundation, the hundreds of millions of people that depend on water from sources replenished by glacial melting, or the rest of us that consume food from crops grown in these areas. There is a growing body of evidence that microbial processes considerably accelerate the melting of ice surfaces, so our work plays a small but important part in understanding how ice melt will change our lives.

Core-firn
5. How has MO BIO been helpful?

I’ve been using MO BIO kits for extracting nucleic acid from environmental matrices for over a decade. I must have extracted thousands of samples. In all that time I have only had six or seven samples fail to yield usable DNA! Soils of all kinds, sediments, biofilms, river water, ice melt, snow, air samples. Even cryoconite, which is particularly challenging as while its biomass is relatively low, it is enriched in humic substances that inhibit PCR. MO BIO kits have made environmental genomics much more accessible.

Samples
Secondly, we have increasingly been facing a challenge I refer to as the “rime of the modern glacier biologist” – ice, ice everywhere, and not a lump to freeze! We work in remote environments, far from the nearest ultrafreezer, and liquid nitrogen or dry ice are impractical to use in deep field. Nevertheless, high quality nucleic acids that are representative of the microbial community at the time of sampling are important to us. Stabilizing solutions such as Soil Lifeguard help a lot.

Arwyn, are you kidding me?  Our Planet Earth thanks you,  your science, and your enthusiasm!  Oh, and congrats on your recent Microbiome Awards win, may your rock star science continue!

 

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Working with Salty Soil

Oct 01, 2014
Michelle Tetreault Carlson
Badwater-Storm-1

Badwater Basin Salt Flats, Nevada, USA

Dear MO BIO,

I am using the RNA Power Soil Total RNA Isolation kit with some high saline soil.   I was not able to extract any RNA but was able to extract DNA with the kit.  What’s going on?

Thao

Dear Thao,

Soils or other samples with high salt content can be a little tricky.  The RNA PowerSoil Total RNA Isolation kit makes use of an anion exchange resin to bind and wash nucleic acids.   Under low salt conditions, the negatively charged RNA/DNA are attracted to the positively charged resin and stick.   Under high salt conditions these charges get screened from each other.  By increasing the salt concentration of the buffer on the column, one can control when the RNA and DNA come off the resin.  RNA will come off first and then with increasing salt, DNA will come off.  However, if you start with a high salt content soil you’ll skew the buffer you run over the resin.   This is what happened in your situation.  The residual salt from the soil was high enough to knock off the RNA but not the DNA.

In order to remove the interfering salt from the soil matrix you can wash it.  Add your sample to a sterile 15 ml Collection Tube and then add up to 10 ml of sterile Phosphate Buffered Saline (PBS). Vortex briefly to mix.  Then centrifuge at 10,000 x g for 2 minutes to pellet the soil and microbes.  Remove the liquid.  You may need to do this more than once, repeating up to three times.  However, it seems to work well.

Best,

MO BIO Technical

p.s.

Soils with high levels of metals can also interfere with MO BIO Isolation kits.  For these types of soils we recommend prewashing with a sterile TE buffer (10 mM Tris, 1 mM EDTA).  The EDTA acts as a chelator.   Like above, you can add the TE to a collection tube with your sample and vortex to mix.  Centrifuge to pellet the cells and soil particles and remove the liquid which will contain a higher fraction of metals.  Repeat as needed.

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Wonderful World of Microbes

Sep 08, 2014
Emelia

Enid Gonzalez-OrtaHello!  My name is Enid Gonzalez-Orta and I am an Associate Professor of Biological Sciences at California State University, Sacramento, where I teach undergraduate students about the wonderful world of microbes.  In addition, I embed an authentic research experience into my microbial diversity class each spring where we study soil samples collected from CSU Sacramento Arboretum or from local vernal pool sites.  We study the bacterial community of these samples using traditional methods, like culturing samples onto laboratory media, sequencing of the 16s rRNA gene through the Sanger method, and building phylogenetic trees.  However, I began to think about how much the field of microbial ecology has changed and is changing.  I thought about how culture-independent methods allow us to peek into environments seldom studied in the laboratory and how these methods reveal members of bacterial communities that were previously not known to inhabit these niches.  I also thought about how next generation sequencing (NGS) methods are becoming becoming the “tradition” in this field.  And, I thought about how important it is for undergraduate students to have hands-on experience with bioinformatics and computing in order to interpret the volume of sequencing data that is produced through NGS.  But, how I could I do this when I myself had little-to-no experience with processing 16s rRNA NGS data?  Then came EDAMAME to the rescue!

 

When the opportunity to attend EDAMAME presented itself, I just couldn’t pass it up.  I haven’t been a student for a little while, so I was excited to be in a classroom sitting in a desk and not in the front of the room for a change.  Not only would this course help me teach undergraduate students in my courses how to process NGS data, but it would help to advance my research program CSU Sacramento as well.   My research focuses on the bacterial diversity of the California Vernal Pool Ecosystem in the Sacramento Valley.   This project is done in collaboration with my fellow colleague Dr. Jamie Kneitel and  has been worked on by many undergraduate students.   One of them, Dana Carper, joined my lab as a graduate student focused her Master’s work on this topic .

 

My experience with EDAMAME so far has been awesome.  We are officially half-way through the course and I can say that I have learned a lot from Ashley, Tracy, and Josh.  And, on top of some great instruction, we have amazing guest lectures!  I can’t wait to take what I’ve learned and teach it to my undergraduates.

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Seagrass: The Whale of the Plant World

Sep 02, 2014

My name is Cassie Ettinger and I am a 2nd year PhD student in Jonathan Eisen’s laboratory at the University of California, Davis. I am working with Hannah Holland-Moritz, a Junior Specialist, and Jenna Morgan-Lang, the Post-doc in charge of the project, to try to characterize the Seagrass Microbiome, the entire microbial community associated with seagrass.

Seagrass

Why study the seagrass microbiome? Seagrass has an interesting evolutionary history.  Starting out as marine algae, an ancestor of seagrass made the transition from marine to terrestrial with its descendants eventually becoming what we know today as plants.  Some of these plants later returned to the marine home world from whence their ancestors had originated (just like whales evolved from a land-based ancestor that returned to the marine environment!). Seagrasses are the only known flowering plants to have made this return to the sea. This is because surviving in a marine environment poses significant physiological and morphological challenges. We are interested in the role microbes might have played in the adaptation of seagrass to the marine environment and if any co-evolution has occurred between seagrass and its microbiome.

Additionally, seagrass plays a vital role in coastal marine ecosystems by cycling nutrients, protecting the coastline from erosion and serving as a food source and hatchery for many marine macrofauna. Microbes have been found to have a significant effect on terrestrial plant fitness, so it only makes sense to try to determine what role microbes might have on marine plant fitness as well.

I am currently investigating the nature of “edge effects” in terms of microbial diversity by analyzing a dataset we have from microbial communities obtained from the inside, on the edge and outside seagrass beds. Hannah is working on determining the intraplant microbial biogeography (how the microbial community composition differs across a single plant) and Jenna is coordinating our collaboration with the ZEN (Zostera Experimental Network), a group of 40+ labs that work on Zostera marina, an abundant seagrass species. They have all agreed to send us leaf, root, sediment and water samples from their respective study sites around the globe! (And of course we are using MoBio PowerSoil kits for all our DNA extractions!)

My goal for EDAMAME is that I will leave the workshop with a better understanding of the data analysis algorithms I am using; I hope that what I learn at the workshop will help inform my choice of parameters and programs to use for my respective datasets. I am super excited to learn more about how to analyze metagenomic data sets so that I can use those skills to investigate the functional roles of different microbes in the seagrass microbiome. I am also enthusiastic to learn how to use R to interpret data obtained from QIIME and mothur and to make beautiful publication ready graphs. I also plan on sharing everything I learn with my fellow lab members back home!

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Dandelions, Microbes, and 300 Middle Schoolers..Oh My!

Aug 25, 2014
Emelia

My email dings soon after lunch on Monday, I open the window, and there it is in all its glory. The MiSeq data has arrived! I promptly download the file giddy with excitement…and wait. And wait. The file is huge, and incomprehensible, and somehow contains nearly 15 million sequences.

 

Luckily, I had signed up to attend the Explorations in Data Analysis for Metagenomic Advances in Microbial Ecology (EDAMAME) workshop hosted by Michigan State University. So here I am at Kellogg Biological Station on Gull Lake taking baby step-by-baby step through the shell, QIIME, Mothur, and R. Many of these I have used in the past, believing that I had taught myself how to run them. But here’s the difference: what I had done was copy and paste some commands, hope it worked, and then compared my results to results from other programs. Very efficient! Now: we are really learning the meat of these programs down to the last –i and / and the knowledge of these details allows the understanding of what happens in each step. My goals while here at EDAMAME 2014 are not to merely analyze this dataset, but to be able to take this knowledge and power and translate it to any dataset.

Stevens

Dandelion photo credit: Lea Shell of yourwildlife.org

 

But, I have gotten ahead of myself. Where did these data come from you ask? The short answer: middle school teachers. The long answer: a bit more complex. I am a postdoctoral research associate at the North Carolina Museum of Natural Sciences (NCMNS) in the Genomics and Microbiology Lab, and, together with the Rob Dunn Group at North Carolina State University, we are embarking on a journey to bring real science into middle school classrooms. Students Discover (http://education.yourwildlife.org) is a program at the intersection of research, education, and citizen science. This summer we had 12 middle school teachers working in the labs at NCMNS, and specifically 3 teachers working with me in the G&M lab on the dandelion rhizosphere. In 3 weeks, these teachers went from learning how to use a pipette, to running an experiment, and finally setting up a multiplexed Illumina MiSeq run. Their excitement and hunger to learn was infectious.

Photo credit:  Lea Shell of yourwildlife.org

Why dandelions? Well first, they are really cool. But importantly: they are everywhere, everyone can recognize them, and they can withstand a lot of disturbance. Perfect for citizen science! So this summer the three North Carolina middle school teachers, Laura Cochrane, Arthina Blanchard, and Amy Lawson, and I transplanted dandelions into nine different soil types to determine what happens to the rhizosphere microbes. This served as a pilot study that will be repeated in their classrooms throughout the school year. Roughly 300 middle school students will be driving research based on the results of this preliminary dataset. By maintaining this connection between NCMNS, NC State, and the classrooms, the students will have an active, hands-on role in current science.

 

For now, building confidence in proper management, handling, and analysis of these data will help foster confidence to move forward in this scientific adventure with some dandelions, their microbes, and a whole lot of middle schoolers.

-Julia L. Stevens

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EDAMAME, onward!

 

EDAMAMEGreetings, Culture Dish followers!

 

My name is Ashley Shade and I am an assistant professor at Michigan State University in the Department of Microbiology and Molecular Genetics. I am excited to share our first days’ adventures in our new workshop, Explorations in Data Analyses for Metagenomic Advances in Microbial Ecology (EDAMAME). EDAMAME is a week-long intensive training in bioinformatics tools and ecological analyses of microbial metagenomics data, held at Michigan State’s Kellogg Biological Station.

 

I know what you are thinking: Silly new professor! Why dedicate preparation time and a week of your life to a non-credited summer course that won’t count towards your teaching requirements?

 

The answer is simple. EDAMAME is happening because there is a hunger for this training. I am leading EDAMAME because I am passionate about informed analyses of metagenomic data. I want to empower other microbiologists to own their analyses, from raw sequencing reads to ecological interpretation.

Edamame

EDAMAME was designed to advance computational skills in students with strong backgrounds in biology. My co-instructors, Joshua Herr and Tracy Teal, both of MSU, and I have worked hard to prepare lectures and hands-on tutorials, and to invite interesting guest speakers. Today is Day 2, and it is amazing what we have already accomplished. Yesterday we dug right into a tutorial on shell. Then, we spent the afternoon yesterday and full day today having the “QIIME” of our life. Titus Brown and Jack Gilbert gave guest research lectures today, and Pat Schloss is arriving tomorrow to work through a day of “mothur”-ing .

 

Did I mention, it is only Day 2?

 

Last evening, the students gave 5-minute lightning chalk talks to introduce themselves and their research. I was impressed by their diverse expertise; their research questions addressed topics ranging from animal nutrition to soil processes. We have students, post-docs, and professors learning side by side. They are tenacious! I like them. I hope that you will be hearing from them with some guests posts in the very near future. (They were happy to have the new MO BIO t-shirts. Photo op to come!)

Shade

If you’d like to join in on the conversation, we are tweeting! #edamame2014. We also have all of our course materials online at edamame-course.org.

 

EDAMAME, onward!

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Where in the World: Salt Marsh Microbes

Want to play in the mud?  Like the ocean?  If your answer to both questions is “yes,” you may have a tinge of jealousy when it comes to this Where in the World project.  François Thomas, PhD, works as a postdoctoral scientist with Stefan Sievert, PhD, at Woods Hole Oceanographic Institution on Cape Cod in Massachusetts.  His collaborative project* funded by the National Sciences Foundation studies microbes in New England salt marshes. By sampling sediments and roots of the salt marsh grass known as Spartina, he uses both cultivation and molecular techniques to understand the microbial communities in each environmental niche. In particular, he is studying sulfur-oxidizing microbes, which can profit from the release of oxygen from the roots and might play a critical role in linking the carbon, nitrogen and sulfur cycles. To get back to your childhood memories, check out this video from the field.

*Anne Giblin, PhD, and Zoe Cardon, PhD, of Marine Biological Laboratory in Woods Hole.

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Troubleshooting Low DNA yields

Jun 24, 2014
Michelle Tetreault Carlson

One of the top five technical concerns we get at MO BIO relates to low yields of DNA.   Whether it be sediment, swabs, waste water or sludge, at some point we’ve probably heard that you didn’t get enough DNA out of it.  After all, there is no such a thing as getting too much DNA.   Nope, haven’t heard that grievance even once!

mb110dWhen someone tells us they didn’t get enough DNA, we rule out the obvious offenders. It could be that your lab-mate left your samples out all night at room temperature and didn’t tell you. It might also be that you’ve made an error in the way you’ve interpreted or conducted a protocol or perhaps you’re using an inappropriate kit. The list goes on and on.  However, the answer could be a lot simpler.  Low DNA yields (defined here as those that can’t be easily measured with an absorbance spectrophotometer, for example, a Nanodrop) may not indicate that something went wrong at all.   It might just be that there wasn’t a lot of DNA to begin with.  This is very common with low biomass samples like swabs, filters or sediment.

Whenever you’re working with a sample that has never been tested, you may not know the taxa or number of cells present, so you may not know how much DNA to expect.   This is especially true of environmental samples.  For example, every type of soil is different, with varying pH, humics, carbon content, etc.   But fortunately you don’t need to march blindly into the darkness.   Instead you can do something much better;  run a control.

Soil in bead tubeFor soil, a good control is to “spike” your sample with a known quantity of bacteria or fungus and run it through the kit protocol.   You can start by growing an overnight culture of your favorite bug.   It’s best to pick something that is representative of what you might be looking for in your sample.  Then, take 1-2 ml of the culture and centrifuge to pellet the cells.  A good rule of thumb for a bacteria like E. coli is that 1 ml of an overnight culture contains 10^9 cells or approximately 5 micrograms of DNA.  Fungus have roughly 10 times the  concentration of DNA per cell as bacteria.   Find out  the genome size of your test bug’s DNA, measure the cell density in your culture and calculate the expected yield.  Now, you can spin down the cells from the culture into a pellet and then resuspend  in a small amount of neutral buffer like 100 uL saline.  Add this to your soil sample and let incubate for at least 10 minutes before starting your protocol.  Extract this alongside your original samples and voila;  you’ll be able to see how much DNA is present in your sample because it will be the difference  in concentration between your spiked and unspiked sample.  If not, then there might be something else going on.   Perhaps there is something in the soil like metals that is interfering with your prep or your homogenization protocol isn’t adequate for cell lysis.  If you can’t figure it out, contact MO BIO Technical support.

Similar “spiking” experiments can be done with other samples as well.   Just treat the cultured sample as you would your working sample.    For example, if you are filtering pond water then you can spike in your microbial culture and filter just the same.    In this way you can be confident that your DNA Isolation method is working properly, determine the concentration of  your sample and eliminate one more scientific unknown.   In fact, it’s always a good idea to run this type of control, even if you’re comfortable with your current extraction method.   It’s a little extra time up front but will likely save you loads of trouble down the road.   Here is to basic science 101!

 

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A Birthday Salute to a Special Sub

Jun 09, 2014
Emelia

Last week marked the deep sea submersible Alvin’s 50th Birthday.  This translates to 50 years of unprecedented ocean exploration and discovery!

MO BIO Laboratories, Inc. would like to thank Alvin and the very dedicated folks that make Alvin’s story so historic in scientific discovery!

Thanks to Chris Linder, Alvin and MO BIO’s cultivating curiosity were united on R/V Atlantis during the first scientific expedition with the newly outfitted Alvin, awesome sauce!  I can’t help but share my own special moments with Alvin, thanks for checking in!

 

GetInline

 

 

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