Erin received her PhD in 2005 from UCLA and during that time focused much of her efforts on building programs for teaching microbiology to students in a way that would engage and fascinate them. Indeed, her CV contains a long list of undergraduate teaching and instructional development activities, curriculum development projects, and education publications. Her dedication to microbiology education continues to be her main focus as a academic coordinator at UCLA. Erin is currently the Director of Initiatives in Life Science Laboratory Education and a co-faculty advisor for the Training of Departmental Teaching Assistants program at UCLA.

Erin’s latest achievement is the upcoming publication of a new text book, a collaboration with Jeff Miller, Distinguished Professor at UCLA.  The book, called:  I, Microbiologist: A Discovery-based Course in Microbial Ecology and Molecular Evolution published by ASM Press, will be available in March.

We interviewed Erin to hear more about the book and her teaching  experiences at UCLA.  Read on to hear her innovative methods for educating the next wave of microbiologists.

Q:  What is “I, Microbiologist”?

A:  This curriculum is based on my research lab courses at UCLA.  I teach a new class each quarter with about 30 students.  The students are split into teams of 4.  Essentially the teams must go out to collect soil samples, identify the organisms present in the samples and using their 16S rRNA sequences build phylogenetic trees.  The course is structured such that no single student can complete the objectives on their own, so they must learn to work as a team to complete their study.  Also, part of the class involves teaching students to keep a lab notebook, writing up their results and solidifying research and experimental concepts. 

Q:  Have your students made any interesting discoveries?

A:  Yes, in fact, stage 2 of the curriculum development will involve setting up an online database to make their results available to the general research community.  One specific example involved a study of soil samples collected 1 mile underneath Terminal Island.  This was part of a feasibility project by the City of Los Angeles and Terralog to break down biosolid waste underneath Terminal Island.  The students had to work out growth conditions for those samples that mirrored the environment they were collected, so anaerobic growth conditions in high temperature and high salt.  They identified a number of bacteria that were metabolically unique and interesting to the biofuels industry.  In fact their results provided the impetus to scale up the experiment with an independent study.

Q:  Has this experiential method of coursework impacted other course curriculum at UCLA?

A:  Yes, a colleague of mine built on a nitrogen fixing organism that we identified in the Mildred E. Mathias Botanical Gardens in a later course and we are now applying for funding to have the bacteria sequenced.  UCLA is strongly considering expanding the concept of discovery based research experiences for undergraduates to other areas of life science research.

Q:  How has this curriculum impacted your students?

A:  I believe it gives the students a more accurate picture of what to expect from a career in research.  Anecdotally I’ve seen some students change course from other career plans to move into a research based career.  Our main goal with these classes is to build critical thinking skills and to teach students how to ask the right questions to determine how well a study was constructed when they are reading a report of the results. 

Q: Which of your own publications was the most influential to you and inspired you to do what you are doing today?

Sanders ER, Karol KG, and McCourt RM (2003). Occurrence of matK in a trnK group II intron in Charophyte green algae and phylogeny of the Characeae. American Journal of Botany 90: 628-633.

The reason is because it reflects the research efforts of my undergraduate years at DePaul University working with two colleagues, both of whom are close friends to this day (Rick McCourt and Ken Karol).  I worked in Rick’s lab for two years and was trained by Ken, who patiently took we undergrads under his wing and taught us every technique and it’s associated nuances.  It was a wonderful experience and inspired me to continue my quest for a career in science.  This experience prepared me for my graduate work — it got me motivated and excited about the world of research.  The fact that I published a paper as a first author undergraduate really underscores the importance of getting students engaged in the process of authentic research early in their college careers — it validates my experience and highlights the potential we all have to thrive and make meaningful contributions to the science community even as students.

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Thanks Erin for your time and especially for your dedication to teaching the critical science skills early on. I am sure that your efforts have a great impact on the students you train and on UCLA’s reputation as a scientific institution.

The book,  I, Microbiologist: A Discovery-based Course in Microbial Ecology and Molecular Evolution will be available in March from ASM Press and can be pre-ordered now. 

If you are an educator or interested in building similar types of programs at your institution, you will want to get a copy of this book.  Also check out www.asmcue.org where Erin will be presenting a poster at the 17th Annual Conference for Undergraduate Educators (May 20-23, 2010) at the Town & Country Resort and Convention Center, San Diego, CA.

Thanks for reading! If you have any questions for Erin or comments about this article, please feel welcome to leave them here on this blog and we will reply as soon as possible.

~Suzanne

 To make scanning the monthly journals a little bit easier for you, today’s blog is a first in what will be a regular series of articles highlighting some of the exciting research going on in microbiology. After scanning the January and February publications on Highwire, I picked a few headlines that I thought the community would find particularly interesting.

There are so many to choose from that I could write all day about the great papers coming out from our customers but I had to narrow it down for the sake of brevity. Most of my picks will lean towards methods papers that may be helpful to our readers, and also research into novel areas of microbiology or “firsts”.  In some cases, papers are chosen to highlight a particular lab group.

These were my top picks covering January and February:

1.  A Rapid and Specific Method for Quantifying Streptomyces Competitive Dynamics in Complex Soil Communities
Daniel C. Schlatter, Deborah A. Samac, Mesfin Tesfaye, and Linda L. Kinkel
Appl. Envir. Microbiol., Jan 2010; 10.1128/AEM.02320-09.

The authors developed a sensitive (down to 0.01 pg of DNA, which is 1 genome equivalent for Streptomyces) and specific qPCR assay for determination of two species of Streptomyces that co-exist but compete with each other in soil. The assay was designed using the short hyper-variable region of the 16S rRNA gene. SYBR Green primers were designed for S. lavendulae and S. scabies using species-specific forward primers and a single conserved reverse primer from the hypervariable region. Because of the background detected from indigenous Streptomyces in soil, a probe based assay was developed using species-specific forward and reverse primers and a Streptomyces specific dual-labeled probe (FAM and NFQ). The specificity of the assay was validated by spiking in S. lavendulae and S. scabies either together or separately into autoclaved soil and in field samples containing 15 different strains of Streptomyces. The assay provides a model for developing qPCR based microbial tracking assays in samples with very complex communities, such as soil

2. Sample pooling masks PCR-based estimates of soil microbial richness and community structure
Daniel K. Manter, Tiffany L. Weir, and Jorge M. Vivanco
Appl. Envir. Microbiol., Feb 2010; 10.1128/AEM.03017-09

Due to the richness of microbial communities in soil, profiling of the  dominant community structure is typically done by taking multiple samples of soil from a plot and combining them to provide a samples that is representative of the soil.  But does pooling of soil or DNA prior to PCR have an impact on the results of community diversity? This paper shows that it does. In this report,  ARISA (atomated ribosomal intergenic spacer analysis) technique was used to analyze the differences in community profiles from three different types of soils processed three different ways. The three strategies were: 1) soil samples were unpooled, extracted, and analyzed individually, 2) DNA extraction from multiple samples were pooled prior to PCR, and 3) soil samples were pooled prior to DNA extraction and then DNA isolated from a large quantity of soil.    Their results showed that pooling soil or DNA prior to PCR results in missed detection of fungal and bacterial phylotypes compared to unpooled samples which revealed a number of amplicons unique to individual samples. Loss of fungal community richness was impacted greater by pooling than bacterial community richness.  This effect differed between sites related to the evenness of the community in the different soils. The authors make some suggestions for best practices in sampling and pooling based on your project goals and soil type.

3. First Autochthonous Case of Rhinocladiella mackenziei Cerebral Abscess Outside the Middle East
Hamid Badali, Jagdish Chander, Shaifali Bansal, Atul Aher, Surendra S. Borkar, Jacques F. Meis, and G. Sybren De Hoog
J. Clin. Microbiol., Feb 2010; 48: 646 – 649.

Cerebral phaeohyphomycosis is a rare infection of a melanized fungi that causes brain abscesses that typically leads to death. The fungi in the order Chaetothyriales, of which Rhinocladiella is a member, are neurotropic in humans, and R. mackenziei causes 100% mortality in even immunocompetent hosts. The organism has never been isolated from the environment and has only been identified patients in the middle east.  This report describes the first case of R. mackenziei infection in a person outside the area of endemicity.  The details of the microscopy and culture techniques used to diagnose the patient are described.

4. Identification of a Two-Component Regulatory Pathway Essential for Mn(II) Oxidation in Pseudomonas putida GB-1
Kati Geszvain and Bradley M. Tebo
Appl. Envir. Microbiol., Feb 2010; 76: 1224 – 1231.

Oxidation of manganese by bacterial species via a family of enzymes known as multicopper oxidases (MCO) serve important potential functions including the binding of trace metals, making Mn oxidizing bacteria good candidates for bioremediation of heavy metal contaminated sites. Pseudomonas putida GB-1 is a Mn oxidizing strain that is easy to work with in the lab and contains an MCO family of enzymes. Their surprising results found that oxidation in this strain was not due to the MCO but involved a sensor histidine kinase (mnxS1) and a response regulator gene (mnxR). Environmental signals, such as nutritional state, growth phase, or oxygen concentration, result in autophosphorylation of mnxS1 and subsequently the transfer of the phosphoryl group to the mnxR transcription factor, inducing downstream effects on genes transcribed by σ⁵⁴ -RNA polymerase dependent promoters.  Their results provide the Mn oxidation field a powerful tool for studying the signals that trigger oxidation.

5. Up-regulation of integrin expression in lung adenocarcinoma cells caused by bacterial infection: in vitro study
Sean Gravelle, Rebecca Barnes, Nicole Hawdon, Lee Shewchuk, Joseph Eibl, Joseph S. Lam, and Marina Ulanova
Innate Immunity, Feb 2010; 16: 14 – 26.

Integrins are heterodimeric transmembrane proteins that interact with extracellular matrix proteins and are important for repair of damaged tissue, regulating inflammatory responses, and tissue remodeling. Integrin receptor expression changes in pulmonary epithelium under pathological conditions, including damage or neoplasia in the lung. One organism that causes severe pulmonary infections in immunocompromised people is Pseudomonas aeruginosa. This study examined whether infection with P. aeruginosa alters integrin expression. Using flow cytometry and RT-qPCR, their results demonstrated a rapid up-regulation of integrins in response to internalization of live, virulent, piliated and LPS containing bacteria. The authors propose a model for integrin signalling in response to infection by P. aeruginosa.

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This small sampling of papers covers a wide range of new and exciting research in microbiology. 

I know there is so much more so I encourage you to leave a comment and let us know what paper or papers came out recently that you think will have a big impact on your field and why. Let us know what are your picks for hot papers in microbiology.

Introduction:

Amoebae exist in the environment in soil and environmental waters, but not in the potable ground well water. Or so we thought.  It turns out that free-living amoeba can and do get into the drinking water system and in fact were responsible for the deaths of two children in Arizona of primary amoebic meningoencephalitis (1).  This was unusual because it was assumed that amoeba do not exist in well water because of the lack of a carbon source and because most wells have mesh casings that prevent passage of species larger than 5 microns in size. N. fowleri is 7-20 microns in size and so are well above the cut off for the well water barrier.

With this knowledge, Ian Laseke et al. (2) investigated the extent of the presence of the pathogenic amoeba N. fowleri in the warm ground water aquifers in the Phoenix area over a range of seasonal collection times and water temperatures. They also looked at different water pumps to see if the oil based pumps influence bacterial and amoeba growth.

Using molecular analysis methods, including an N. fowleri  specific PCR assay (specific for the  Mp2Cl5 gene)  and sequencing of 16S rRNA gene libraries made from the total bacterial DNA in water samples, they were able to identify amoeba positive wells and then make specific and clear correlations between the microbial communities in positive and negative wells.  To see what they discovered, keep reading.

Methods:

Well Selection: Six public water supply wells in Phoenix were sampled between winter of 2004 and autumn of 2005. To determine if presence of N. fowleri was sensitive to water temperature, the selection of sites were based on both the time of year and the water temperature at individual wells . In addition, the pump type (submersible or oil-lubed turbine) was taken into consideration to examine if  contamination of the water with oil lubricant was playing a role as a food source for bacteria that were serving as a food source for the amoeba. Control water negative for amoeba came from the Mason Drinking Water Treatment Plant in Cincinnati, Ohio.

Sampling and Filtration: Each water sample (collected in December, August, and September) were filtered through serially arranged filters (Micro-Wynd II D-PPPY, Cuno Inc.) with pore sizes of 0.5 and 1.0 micron, respectively. Volumes ranged from 236 to 4402 liters. Filters were transported to the lab at -80^oC for processing.

The negative control water was filtered through pore size 0.5 micron catridges and three 100 liter control samples were spiked with 10³, 10⁴, and 10⁵ N. fowleri cells and then filtered through 0.5 micron pore size filters similarly to the uninfected water.

DNA Extraction: A portion of the filter (~1.61 cm² ) was removed using a sterile razor and processed using the PowerSoil DNA Extraction Kit to isolate DNA. DNA was isolated from pure cultures of N. fowleri using the same method and templates were used for PCR.

Molecular Assays:  The PCR assay was a nested PCR with the first round of PCR targeting the Mp2Cl5 gene unique to the genus Naegleria and the second round of PCR was performed using primers targeting a conserved region of the Mp2Cl5 gene specific to N. fowleri (3). PCR reations were analyzed using 2% agarose gels.

To identify the predominant bacteria in the well water, 16S rRNA gene clone libraries were generated from selected samples. Clones were sequenced and phlyogenetic trees were constructed and analyzed.

Results and Discussion:

The nested PCR assay for detection of N. fowleri  was sensitive down to 10 cells/ml of water and using this method, approximately 27% of all samples tested positive N. fowleri.  While all of the samples that tested positive were from the summer and fall months, water temperature ranged from between 29 to 47^oC. So while infection of the water may be seasonal, it is not related to water temperature. It was also not related to the type of pump, indicating that the contamination of oil in the water does not enhance the level of infection of amoeba.

To look for linkages between the bacterial populations in water, which are a carbon source for amoeba, and the presence or absence of N. fowleri, the 16s rRNA gene libraries were analyzed and compared. Wells that were negative for N. fowleri consisted only of β- proteobacteria (primarily clones were within the Caldimonas and Leptothrix clades).  Clones from the positive wells, however, contained a much higher level of bacterial diversity.  N. fowleri positive wells contained members of the Bacteroides and Firmacutes phyla, as well as Deinococcus- Thermus, Nitropira, and Acintobacter.  

Summary:

Interestingly, the presence of a high level of iron and manganese oxidizing bacteria (such as the genera Caldimonas and Leptothrix)  in negative wells correlates with previous findings that amoeba require these metals to be present in environmental waters to thrive (4, 5). It has been noted that amoeba generally prefer gram-negative nonpigmented bacteria as a food source and pigments are known to possess iron chelating activity (6).  All of the negative wells were reported to have very low levels of iron and magnesium. 

To summarize, municipal warm ground water wells in the Phoenix area tested positive for N. fowleri and their presence did not correlate with water temperature, pH,  or heterotropic plate count.  Presence of the amoeba did correlate with a more diverse bacterial community, specifically an increase in some of the enteric bacterial groups and a reduction in metal oxidizing β- proteobacteria. This leads to some exciting future studies on the role of metals and metal-oxidizing bacteria on the amoeba diet.

Final note:

Personally, I would be interested in more studies on what caused some of the ground well water microbial communities to change and how to restore them to an amoeba inhibitory state. The questions that come to my mind are- what happened first; did the amoeba arrive and then the ecology changed? Or did the microbial community change and the amoeba flourished?  In addition, it might be interesting to isolate, if possible, live N. fowleri from positive water wells to determine if their size has diminished to allow them access to water wells or whether the mesh casing meant to keep large organisms out of the water wells needs to be replaced.

Thanks for reading and if you have any questions or comments, please leave us a note below or email us anytime at technical@mobio.com.

For more advice on choosing and handling filter membranes, look at this previous article on water filters. Included is a video on how to handle the water filter without damaging the membrane suface.

Have a great week,
Suzanne

References:

1- Marciano-Cabral, F. et al. 2003
Identification of Naegleria fowleri in domestic water sources by nested PCR
Appl. Environ. Microbiol. 69:5864-5869

2- Laseke, I. et al. 2010
Identification of Naegleria fowleri in Warm Water Ground Water Aquifers
Journal of Environmental Quality, vol. 39, January-February 2010, page 147-153

3- Reveiller, F.L., et al. 2002
Development of a nested PCR assay to detect the pathogenic free-living amoeba Naegleria fowleri.
Parasitol. Res. 88:443-450

4- Duma, R.J. 1981.
Study of pathogenic free-living amoebas in freshwater lakes in Virginia.
Environmental Health Effects Research Series. EPA no. 600/1-800-037

5- Kyle, D.E., and G.P. Noblet. 1985.
Vertical distribution of potentially pathogenic free-living amoebae in freshwater lakes.
J. Eukaryot. Microbiol. 32:99-105

6-Singh, B.N., and G.D. Dutta. 1984.
Small free-living aerobic amoebae: Soil as a suitable habitat, isolation, culture, classification, pathogenicity, epidemiology and chemotherapy.
Indian J. Parasitol. 8:1-23

It’s important to love your work and fortunately for us, there is so much to love about microbiology and the environment. But to find out what is best about working in this field, I asked the question to several of my scientist friends:  What do you love about your work? Why do you study environmental microbiology and what is it that makes it the best field of work?

And below are some of the best responses. Some are my own, but most are responses I received from people who study some of the most unusual samples from the most extreme environments in the world.  I think you will agree that environmental microbiology provides experiences unlike any other field. Let us know your reasons for loving your work!

14 Reasons to Love Environmental Microbiology:

1. You get to play outside in the mud, snow, water or clouds (see picture at end of article).

2. There is virtually an unlimited number of research projects to choose from. “Microbiologist William B. Whitman, estimates the number of bacteria in the world to be five million trillion trillion. That’s a five with 30 zeroes after it. Look at it this way. If each bacterium were a penny, the stack would reach a trillion light years.” (1)

3. Your research will have an impact on everything living on the planet, humans, animals, and plants. Basically all the Kingdoms benefit from what you do.

4. You have the opportunity to visit exotic and remote locations.  

Graduate student Rick Davis explains, 

I think I’ve been really lucky with the places I get to study– I got to go to Samoa, Hawaii, and Yellowstone this year!

He also added reason number 5: 

Environmental microbiologists are more laid back and we are generally more collaborative than competitive, which allows for greater progress and more fun at conferences!

And since I met Rick at the 2009 Gordon Conference for Applied Environmental Microbiology, I have to agree with him!

John Mackay, a molecular biologist and director of business development at the plant diagnostic company, Linnaeus, tells me:

6. You can cruise around the seas for months, sequence a bit of sea water and write the whole lot off on your research grant!

7. You can work on things you can eat or drink – I recommend wine and truffles!

8. When you find new species (almost a given!), you can name them after yourself.

 New discoveries are also what motivates Charlie Lee from the University of Waikato, a Postdoctoral researcher in microbial ecology studying the Dry Valleys of Antarctica. He echoes the sentiment that discovery is almost a guarantee:

9. Most systems we look at are relatively poorly understood, and it’s always exciting to discover something for the first time.

 Tom Niederberger, a Postdoctoral researcher in marine biosciences at the University of Delaware, has more to add:

10. The international travel is a great reward. The world is your playground as microbes have colonized basically all habitats on earth, and it’s great to travel around sampling not only the microbes, but new cultures/food/travel etc. and not being chained to the lab and pipette. Also the international collaborations and conferences also are great. (authors note: I met Tom at the same 2009 Gordon Conference where I met Rick. It is true. The conferences are great.)

11. But I think what is most important is that microbes in the environment are essential not only for the health of the planet (e.g. global nutrient cycling / global climate change) but they are also intimately linked to the healthy functioning of our bodies. i.e. the are really important!

12. Also, as both you and Charlie touched on, is the excitement of the unknown. Most of the organisms cannot be cultured and we know nothing about them…I think this is great motivation and it will keep you busy, and there are always new problems to solve and new questions to ask.

All excellent points!

And from a molecular biologist from Colorado State University (who wished to remain anonymous) come two excellent points I hadn’t considered:

13. Extremists don’t kidnap environmental microbiologists. Actually, they give them back.
 
14. If you get tenure, who’s going to boot you out?  Exxon?

Did I mention that environmental microbiologists are funny?

To drive home the point of how much fun environmental microbiology can be, check out this photo of the students in the lab of Laurie Connell after a day collecting clams in Maine. One of Laurie’s many projects involves looking at populations of mutant clams that can resist red tide toxins.

It certainly looks like it was a productive day at work!

Left to right: Janice Duy, Kendra Waters, Willis Beazley and Amber Bratcher.

Thanks to all for your fun and insightful comments!

Tell us your thoughts about your work. Why do you love what you do? What makes environmental microbiology such a great field to you?

Let us know- send us your comments either on the blog or email us at technical@mobio.com.

Have a great long weekend!
MO BIO will be closed on February 15th in observance of President’s Day.

Suzanne

P.S. Do you want a MO BIO Flag? Are you going on a field trip collecting samples? Take pictures of your excursions and we’ll enter them in our “Where in the World” program where you can earn free t-shirts and products! Click here for directions on how to enter.

References:

1. http://www.sdearthtimes.com/et0998/et0998s8.html

Besides cars and roads, turtle survival is also facing threats from new predators; free-roaming cats and dogs who eat their eggs. 

Endangered turtles = endangered wetlands. Maintaining an environment that protects turtles ensures the ecology of Wisconsin is protected for other species that need the wetland to survive as well.

To this end, enter Sean Murphy, a Masters student at University of Wisconsin-Parkside. Last month, MO BIO Labs had the chance to chat with Sean about his research on wetland conservation and monitoring. Sean explained how the genetic mapping of turtle species within the wetland environment allows them to determine the health of the watershed.

We found Sean’s research fascinating as he told us more about the ways they are using molecular biology to protect and conserve the wetlands of Wisconsin.

Q: How are turtles involved in wetland conservation?

A:  A healthy wetland is integrated with neighboring environments. Wetland environments that are isolated are often an environment that is in distress and in danger of decline. Through population genetics, we can compare the genetics of turtle populations commonly found in wetlands throughout southeast Wisconsin. When there is a steady stream of migration between wetland environments, microsatellite analysis will demonstrate high degrees of heterozygosity in the turtle population. Environments with low heterozygosity or with microsatellites that differ substantially from neighboring turtle populations are likely to be the result of genetic isolation. That genetic isolation may be an indicator of a declining or distressed watershed. This is the focus of our research.

Q: Why focus on turtles as the indicator species?

A: Turtles are relatively sedentary and long-lived. They prefer to stay in one area, however, in active seasons they may travel to other wetlands to nest, often traveling as much as a mile to mate. Additionally, they require a varied environment, with sandy areas to nest and shallow water to hibernate. The annual migrations mean that isolated populations are not likely to naturally occur. Further, in the Great Lakes region the development of wetlands areas have left the Blanding turtle on the endangered list in many states and on the threatened list in even more. So the connection between wetland destruction and the decline in turtle population has been clearly observed.

Q:  How has MO BIO Laboratories and our products assisted you in furthering this work?

A: Our turtle blood samples are particularly clotted and it is very difficult to break apart these clots. MO BIO’s UltraClean Tissue & Cells kit has a bead beating step in addition to a proteinase K digestion which really helped us to ensure that DNA wasn’t being lost when we passed the sample over the spin column. Adding to this complication, turtle blood has nucleated red blood cells so the yield of DNA is extremely high. Again the Tissue & Cells kit overcame challenges with spin column clogging that we experienced with other kits.

*****

Thank you Sean, for sharing with us your passion for environmental conservation. We are glad we can play a role in this important research!

Funding for this project is provided by the Root-Pike Watershed Initiative Network. Additional support comes from Dr. Gregory Mayer, Associate Professor at University of Wisconsin – Parkside and Robert Jagla.

Here is more information on how you can protect wetland turtles and reptiles and the Wetland Turtle Group site gives information on endangered species of turtles.

Answer:  To Get to the Other Wetland!

Have a great week folks!

Suzanne

Problems achieving high yield and purity are exaggerated in environmental samples because of the added complexity of  microorganism lysis and inhibitor removal.  Quantifying the nucleic acids in these samples is the easy part. But if you don’t know what to look for, you can easily make mistakes in interpreting the results, which can lead to a lot of repeat work or missed critical information in your experiments.

Let’s discuss some of the common misconceptions surrounding DNA isolation and quanification and what problems to look out for before going to the next step with your sample.

1. True or False:  A higher UV A₂₆₀ reading means more DNA.

False.  A high A₂₆₀ reading does not always mean high genomic DNA yields.  One of the main reasons for a high A₂₆₀ reading that does not correlate to genomic DNA is the absorbance of UV due to highly degraded DNA or RNA. Degraded RNA absorbs a high level of UV and results in a boost to the A₂₆₀ reading. The method used to purify the DNA after lysis will determine what is present in the final sample. Many purification methods do not separate the small DNA and RNA from the high molecular weight gDNA. The PowerSoil DNA Isolation Kit does.

The only way to know what you really have in your sample is to run 5-10 µl on an agarose gel. This will give you a clear indication as to whether you have predominantly genomic DNA or a mix of nucleic acids sheared to varying lengths.

2. True or False:   Bead beating always gets higher yields of DNA

False. Not all samples need to be homogenized with high velocity bead beaters. Typically for RNA extraction of tissues and plants, definitely use the strongest method available to you. You want to break the genomic DNA down in size. For genomic DNA from a variety of environmental samples, gentle methods such as vortexing with beads will isolate high molecular weight DNA. When measuring the yields on a spec, as mentioned above, the more sheared the DNA, the higher the absorbance reading. This does not mean more DNA was isolated.  When qPCR is performed, less DNA will be added to the reaction due to the false high reading, leading to higher Cq values and inaccurate quantification in the sample.

For microbial DNA from soil, the bead beater will tend to do more damage than good. For DNA from fungus and spores, a thorough discussion on ways to optimize the lysis using different beads and heat has been described in detail. We also give some recommendations for using the bead beater and soil based on published references.

The best approach to ensure the integrity of the DNA is to run a gel in addition to the Nanodrop or UV scan so you can make a better assessment of what you really have. If you see a smear on your gel, then the bead beating was too hard.

3. True or False:  If the sample looks clear, it is free of humic and fulvic acids

False. Humic acids give the sample the characteristic brown color so if your DNA elutes with color, you know you’ve got a lot of contamination. Even if it looks clear, there can still be low levels of humic and fulvic acid or even polysaccharide contamination in the sample. Using a kit with Inhibitor Removal Technology such as PowerSoil, PowerWater, and coming soon, PowerBiofilm, will ensure that a clear eluate is actually clean.

In addition to the A₂₆₀ reading for yields, a low 260/230 ratio can be indicative that the sample still has some organic contaminants. A ratio above 1.5 is ideal. A ratio below 1.0 has significant contaminants present that may interfere with enzymatic applications.

4. True or False: If my soil has low amounts of DNA, I need to try stronger methods for lysis

False. If your soil has low amounts of DNA, you need to start with more.

The yield in soil varies greatly but for a rich organic soil with a high microbial load, yields of DNA will range between 20-30 µg per gram of soil (5-7 µg/ PowerSoil prep). According to Whitman et. al (1998),  rich top soil contains 1-2×10⁹cells/gram of soil (1).  Using an E.coli genome as an example, this equates to 5-10 µg of DNA in a gram of soil or around 1-3 µg of DNA per PowerSoil DNA Kit prep. Eluted in 50 ul, the concentration to expect for microbial rich soil is around 20-60 ng/µl. If the genome of the organisms in soil are double the size of E.coli, then yields of 40-120 ng/µl are in the correct range for microbe dense soil.

Most of the soil we use in our lab are not at the high end of microbial load, so yields of 10-20 ng/ul are not uncommon for average soil preps.

Given this information, if another isolation method gives you yields far above this range, it is not all DNA. The DNA is either contaminated with UV absorbing PCR inhibitors or mostly degraded RNA. Either way, the information obtained from genotyping will not be as complete or accurate as clean pure microbial DNA obtained from using PowerSoil and UltraClean Soil Kits.

Make sure to always check the yields on a gel and for even greater accuracy, use qPCR.

 5. True or False:  PCR is the best way to check for inhibition

True. The only way to know if the sample is inhibitor free is to use it in an enzymatic reaction. Even better is to use qPCR and perform serial 10 fold dilutions and check the efficiency of amplification using a primer pair for 16s rDNA. It is likely that there will be some background amplification in the water control because most PCR mixes have background bacterial DNA, but the difference in Cq value between the samples and the control will be far away enough to not matter (usually 6-10 cycles).

With qPCR, the desired result is a change of ~3.3 cycles between each 10-fold dilution. This indicates perfect doubling each cycle and is a sign that the sample is inhibitor free. What you may see is the first sample (undiluted) is shifted to the right and then the rest of the samples fall into place. This indicates that there is some inhibiting substances in the DNA. Remember not to add too much DNA. 1-2 µl per 50 ul PCR is adequate or around 10-100ng for the first sample and then dilute from there.  If the first sample amplifies to soon (and falls in the baseline for the instrument, where fluorescence is subtracted out as background), it will cause some problems with the standard curve so you may want to start with 10 ng and dilute from there.

Another good approach is to set up a PCR reaction that always works (such as for a plasmid) and then spike in 1 µl of the DNA from the environmental sample. If it causes the PCR to fail, or reduces the amount of product, it indicates inhibition.

Summary

These technical tips do apply to more than just soil and water samples. Even DNA from blood or tissues can be affected by inhibitors causing problems with absorbance readings and inaccuracies in quantification. The best approach is to always run a quick agarose gel to go with your Nanodrop results so you can see the integrity and composition of the sample along with the yield. Additionally, PCR or qPCR can help get a more exact quantification of the amount of gDNA or number of microbes in the starting sample.

Thanks for reading, and as always, send us your questions and comments either here on the blog or at technical@mobio.com.

Have a great week!

Suzanne

References:

1. Prokaryotes: the unseen majority.
Proc Natl Acad Sci U S A.
1998 Jun 9;95(12):6578-83.

Next, the sample needs to be cleaned before final purification. This is done using the Inhibitor Removal Technology (IRT) process. Continuing our article at the second stage of the protocol, inhibitor removal, let’s go into greater detail on the different aspects of the chemistry used for the purification of DNA from soil using silica spin filters.

Removal of Inhibitors:

We are now ready to remove the PCR inhibiting substances from the soil homogenate. The humic acids are what give the sample the brown color. Present will also be polysaccharide if your soil sample had plant material or even some biolfilm content. Removal of inhibitors is what makes MO BIO and the PowerSoil kits stand out. MO BIO Labs developed a patented method, called inhibitor removal technology (IRT) to precipitate out the humics, phenolics, and polysaccharides from lysates.  IRT involves a two-step process where-by the proteins and debris are removed first followed by flocculation of large insoluble macromolecules. After using inhibitor removal solution (IRS), samples typically look clear. The PowerSoil and PowerWater protocols are all optimized for the amount of IRS needed to clear even the most problematic samples, however, more can be used if inhibitors are still present (as determined by PCR).  Repeat flocculations with IRS are ok to do as necessary.

The IRT steps are performed using cold temperatures to enhance the flocculation but for the second step (IRS), we recommend not to extend the time significantly over five minutes. Lower DNA yields may result from prolonged incubation in IRS.

Stopping points?

If you need a stopping point in the PowerSoil protocol, the best place to pause is after the IRS step and before adding the binding solution. The lysate can be frozen at -20°C and used for binding to silica spin filters the next day.

Binding to Silica Filter Membranes:

At this point, the DNA is ready for purification on a silica membrane. The lysate should look clear (can be slightly yellowish if the soil was heavy in organics). For the DNA to be captured on silica membranes, it requires the presence of a high level of chaotropic salts. The ratio of the binding solution (Solution C4) to the lysate is critical for good yields. If too much is used, recovery of degraded RNA will result. If too little is used, a portion of the high molecular weight genomic DNA is lost. For this reason, we instruct you to take up to 750 µl of your lysate into this step so that the entire lysate will fit in the 2 ml collection tube once the 1.2 ml of binding salts are added.   

If you need to take more than 750 µl, you will need to increase the binding solution as well. A good ratio is two volumes of binding solution C4 per sample volume. You will need to split the sample into two 2 ml collection tubes or a larger tube (5 ml or 15 ml) to make sure everything is well mixed.

Vacuum Manifold Option:

Normally, if you followed the standard protocol, binding to the spin filter requires three loadings of the column. One way to speed this process up is to try the PowerVac Manifold System.  It is very fast and easy and results in less handling. If you have a vacuum manifold already, then all you need are the PowerVac Mini Spin Filter Adapters. In our lab, we regularly use this method to speed up processing.  If you decided to use more of the lysate than recommended and increased the amount of binding salts, using the vacuum manifold will be the best way to reduce the time required for loading the column 4 or 5 times.

Washing the DNA:

Because of IRT, most of the soil related contaminants are removed so the column will not need a heavy salt wash like with other kits. The washing step here is needed to remove the chaotropic salts from the column. If any salt is left behind on the column membrane, the DNA will not elute efficiently and the DNA that does elute will be contaminated with guanidine. To remove salts from the column, the wash buffer contains ethanol which solubilizes and rinses away salt. One wash typically does the trick. However, if you are having problems with low 260/230 readings (as observed by high 230 absorbance on a Nanodrop), then a second wash may be performed. If you run out of wash buffer, 100% ethanol can also be used to wash the membrane as well. We use 100% ethanol on the vacuum manifold protocol and this can be used manually in the event you run out of wash solution and need more.

Don’t forget to spin dry the column before elution so the DNA can be eluted efficiently. Left over ethanol on the column will make the DNA release from the membrane inefficient.

Elution:

The final step is releasing your DNA from the membrane into a 10 mM Tris pH 8.0 buffer.  DNA dissolves faster in a neutral to slightly basic pH. You may use water to elute but because water tends to have a low pH (usually around 4-5), the efficiency could be reduced. One hint for an increased yield during elution is to allow the buffer to incubate on the membrane a few minutes at room temperature before centrifugation. Incubation from 1-5 minutes will help resolubilize the DNA in a smaller volume. Don’t elute in less than 50 µl or you will leave too much DNA behind. 

Your DNA is now ready to use in PCR or for gel electrophoresis!

FAQs:

How much DNA is typically in soil?

After all of this discussion, you may be wondering how much DNA can I expect from soil? The answer is that it varies. The moisture content, organic content, and where collected will all play a role.

In our labs using “normal” soils or temperate soils, such as garden soil, the yields can range from 2-5 ug of DNA per 0.25 gram (a prep). We have worked with some agricultural soils, such as soil from the Strawberry Fields in Carlsbad, and these yields are far lower- around 0.25 ug per 0.25 gram of soil. Sandy and clay soils tend to have lower yields and very low organic content.

What can I do to increase yields in clay and sandy soils?

One current theory with sandy soils and clay soils is that the released nucleic acids are tightly binding to the soil itself. There are several references looking at ways to pre-block soils to prevent loss of the microbial DNA, including the use of skim milk (1). Some evidence suggests that divalent cations are playing a role in DNA binding to the surface of soils (2). For this reason, some of our customers have found success by adding EDTA into the bead tube during the homogenization step at a final concentration of 50 mM.

Summary:

This covers all of our tips and tricks for isolating DNA from soil. To summarize, soils vary widely in their characteristics and microbial load so expect the yields to vary when extracting different samples.   Two key steps for obtaining high yields and integroty of DNA are the homogenization step and the binding step . If your yields are lower than expected, optimization is usually done at these steps.  And remember, using more soil will not result in more DNA.

We are always interested in hearing how you tweaked the MO BIO kits for better results with your samples. Drop us a comment or email (technical@mobio.com) and let us know your special tips and tricks for isolating DNA from soil.

Thanks for visiting!

References:

1. Microbes and Environments
Vol. 19 (2004) , No. 1 pp.13-19
An Improved DNA Extraction Method Using Skim Milk from Soils That Strongly Adsorb DNA
Yuko Takada-Hoshino and Naoyuki Matsumoto

2. FEMS Microbiology Letters
Volume 97 Issue 1-2, Pages 31 - 39
Adsorption of DNA on clay minerals: protection against DNaseI and influence on gene transfer
Eric Paget, Lucile Jocteur Monrozier, and Pascal Simonet

Because of the amount of detail I want to share with you, this will be a two part blog. The first part will focus on the steps involved in homogenization, that being the beads, the homogenization equipment, and the lysis buffer. In the second part of the article, we will focus on the role of the chemistry in achieving optimal binding, washing, and eluting clean DNA.

MO BIO Laboratories has several products for DNA isolation from soil. For the purposes of this article, we will focus on the PowerSoil DNA Isolation Kit because it is our most popular product for this purpose. It uses our patented Inhibitor Removal Technology^® (IRT) for the removal of humic substances and polysaccharides and is performed using a mini spin filter and a microcentrifuge. 

Important notes before starting…

Something to keep in mind is that all soils vary in microbial load and organic content so DNA yield among different soils can vary. Yield is not based on the amount of material processed alone. Even soils collected from the same core but at different depths within the ground will have variable load and organic makeup.

For better consistency…

Consistency in yields between preps is difficult to achieve with soil because each scoop can contain different amounts of organic material, such as plant leaves or debris, insects, pebbles or sand. At MO BIO Laboratories, we sieve the soil for the best consistency so that the texture is uniform and the large particles are removed. If uniform yield among your preps is important to you, sieve first.

More is not always better….

It is important to note, processing more soil does not always yield more DNA. This is because the lysis buffer will be absorbed by the bead solution making sample homogenization inefficient.  Scale up of soil is possible but is soil-type dependant.  The PowerSoil DNA Kit is meant for small scale preps. If you need to process more soil than 0.25 grams, MO BIO offers alternative kits such as the PowerMax^® Soil DNA Kit for 10 grams of sample and the RNA PowerSoil^® Kit with DNA Elution Accessory Kit for starting with 2 grams of soil.

Now let’s go over the DNA Isolation protocol step by step. We will talk first about lysis and specifically about the mechanical aspects of lysis.

Step One: Lysis

High yields of high quality intact DNA requires a strong lysis. The lysis needs to be strong enough to break open microbes and fungus without severely shearing the DNA. There has to be a balance between the types of beads used and the amount of time used for mechanical homogenization. Temperature can also be used to boost lysis of tough organsisms or spores in combination with bead beating.

Bead types: 

There are many bead choices for the lysis of microbes in soil. The type of bead, shape, and size will all impact the DNA yield and integrity.

MO BIO prefers to use a garnet rock type of bead that varies in size and has sharp edges. Because of the size variation, the rocks can help break down both large clumps of soil and grind microorganisms that shake loose. Garnet is soft so will break down into smaller pieces when used in a high powered bead beating instrument. This works fine as many of our customers use them in the FastPrep or Precellys when they want to increase the lysis power for isolation of DNA from fungus (references below).

Other beads may be used, such as 0.5 mm glass or 0.1 mm glass if bead beating in a high powered instrument for longer periods of time is desired. These beads will cause more DNA damage but for very tough organisms, such as spores, it can be helpful. You can even mix the glass and garnet together if you need a combination of large and small

The homogenization equipment:

As described in the previous article, the vortex homogenization method allows for the best integrity DNA and is also the least expensive method. The time for vortex is ten minutes and this gives optimal results for lysis of bacterial cells in 0.25 grams of soil in our lysis buffer. Longer vortex times do not seem to increase the yield and will cause more DNA shearing.

The use of a Precellys or FastPrep is an option if you have one and want stronger lysis. Most customers use these for only 45 seconds to 1 minute for isolation of bacterial and fungal DNA at a setting of 5 on the FastPrep. A setting of 5 m/s on the FastPrep is equal to about 5200 rpm on the Precellys. Some customers prefer using a FastPrep setting of 4 (Precellys setting of 5000 rpm) for 15-30 second intervals and 3 or 4 pulses per sample. As you can see, using a high powered bead beater will require some evaluation on your part to determine the best setting for your sample. A list of references where the FastPrep was used in combination with the MO BIO UltraClean Soil or PowerSoil Kits is at the end of this article.

The lysis buffer: 

The other key ingredient in a strong lysis is the solution used to pop the cells. This buffer needs to fulfill several functions when it comes to soil. First it needs to disrupt cell membranes in combination with the mechanical homogenization. Second, it needs to be gentle enough to not denature the DNA, and third, it needs to work regardless of the pH of the soil. Soils that are acidic need to be neutralized for optimal DNA yields since the acidic conditions are harmful to the DNA. The lysis buffer in combination with the Solution C1 or S1 (in PowerSoil and UltraClean Soil kits, respectively) provides the optimal conditions for microorganism lysis from any soil type.

What about heating?

For those cases where a stronger lysis is desired, besides trying a high powered beating method and glass beads, it can be helpful to heat the sample before beating. An incubation of the soil in the lysis buffer at 65°C-70°C for 10-15 minutes will help to weaken the cell walls before homogenization. This treatment has been effective for spores and fungus.

Another method is to perform freeze/thaw cycles (3) with the soil, alternating between -20°C or -80°C and 37°C. This can enhance cell breakage as well, although might be less convenient than simply heating as described above.

Summary:

OK, we will end here because this is a lot of information. To summarize so far, the lysis step is the area where the most optimization is possible and depending on what you want to do with your DNA, you can go as easy or hard as you need. It is the combination of the beads + the equipment + the buffer that works together to provide you optimal yields and integrity of DNA. Really, this applies to any sample you are lysing whether it is animal tissues for RNA, bacterial cultures for DNA, or biofilm for RNA or DNA.

Fortunately, MO BIO labs R&D scientists are spending a lot of time working out these optimal conditions for a host of environmental samples, saving you time for your experiments.

But we can’t work with every sample type so we would love to hear from you and how you optimized the lysis of your sample type for the best result. Let us know what you do; bead type, time and homogenization method, and which MO BIO Kit you use to get optimal yields of DNA or RNA.

Next week we’ll continue our discussion on DNA isolation from soil and focus on the steps involved in the purification from the silica spin filters.

Thanks for reading!

References for using  high velocity bead beaters and the MO BIO Soil Kits:

Shifts in Microbial Community Composition and Physiological Profiles across a Gradient of Induced Soil DegradationGuilherme M. Chaer, Marcelo F. Fernandes, David D. Myrold, and Peter J. Bottomley Soil Sci. Soc. Am. J., Jun 2009; 73: 1327 – 1334.

Variations in Archaeal and Bacterial Diversity Associated with the Sulfate-Methane Transition Zone in Continental Margin Sediments (Santa Barbara Basin, California)Benjamin K. Harrison, Husen Zhang, Will Berelson, and Victoria J. OrphanAppl. Envir. Microbiol., Mar 2009; 75: 1487 – 1499.

Diversity of Basidiomycetes in Michigan Agricultural SoilMichael D. J. Lynch and R. Greg ThornAppl. Envir. Microbiol., Nov 2006; 72: 7050 – 7056.

Community Structure in the Sediment of a Freshwater Stream with Variable Seasonal FlowSteven A. Wakelin, Matt J. Colloff, and Rai S. KookanaAppl. Envir. Microbiol., May 2008; 74: 2659 – 2668.

Changes in Bacterial and Archaeal Community Structure and Functional Diversity along a Geochemically Variable Soil ProfileColleen M. Hansel, Scott Fendorf, Phillip M. Jardine, and Christopher A. FrancisAppl. Envir. Microbiol., Mar 2008; 74: 1620 – 1633.

Molecular Profiling of Rhizosphere Microbial Communities Associated with Healthy and Diseased Black Spruce (Picea mariana) Seedlings Grown in a NurseryM. Filion, R. C. Hamelin, L. Bernier, and M. St-ArnaudAppl. Envir. Microbiol., Jun 2004; 70: 3541 – 3551.http://aem.asm.org/cgi/reprint/70/6/3541

Mycobacterium aviumsubsp. paratuberculosis in the Catchment Area and Water of the River Taff in South Wales, United Kingdom, and Its Potential Relationship to Clustering of Crohn’s Disease Cases in the City of CardiffR. W. Pickup, G. Rhodes, S. Arnott, K. Sidi-Boumedine, T. J. Bull, A. Weightman, M. Hurley, and J. Hermon-TaylorAppl. Envir. Microbiol., Apr 2005; 71: 2130 – 2139.http://aem.asm.org/cgi/reprint/71/4/2130

Molecular Fingerprinting of the Fecal Microbiota of Children Raised According to Different LifestylesJohan Dicksved, Helen Flöistrup, Anna Bergström, Magnus Rosenquist, Göran Pershagen, Annika Scheynius, Stefan Roos, Johan S. Alm, Lars Engstrand, Charlotte Braun-Fahrländer, Erika von Mutius, and Janet K. Jansson Appl. Envir. Microbiol., Apr 2007; 73: 2284 – 2289.http://aem.asm.org/cgi/reprint/73/7/2284

Welcome back to The Culture Dish! We hope you are all having a great start to your week and to the year 2010.

We are excited to begin writing articles on a whole range of topics all aimed at helping you with your research.

I wanted to begin by letting you know about a promotion we’ve been running since December just for our blog readers.  The promotion is called Tea for Two:

Leave a comment on any of the articles posted on the blog and on February 1st, one reader will be chosen to win a free  25 pack Ethidium Bromide Tea Bags and a $5 gift card for Starbucks for a coffee or tea. That’s tea for your science and tea for you!

So don’t be shy. We want to hear what you think.

Countdown to ASM!

We also wanted to remind you that abstracts for 110th Annual Meeting for the American Society of Microbiology are due January 13th. ASM is one of the most comprehensive and highly regarded microbiology and life science conferences in the world. Come to beautiful San Diego for ASM from May 23rd-27th and meet our team.

 This year, make sure to mention MO BIO’s name and the kit you used in your poster or talk methods section and let us know your poster day and number once you have it. We will be giving researchers who cite MO BIO extra attention. More details will follow in the newsletters and blogs in the coming months.

We are really looking forward to meeting you in 2010!!

For the final blog for 2009, we wanted to give our readers an idea what a typical day in R&D at MO BIO was like this past year.  To do that we thought it would be fun to share with you 10 phrases you might have heard if you were walking past the R&D labs anytime during the day.  All of these comments relate to products we launched throughout the year or samples we worked on to help our customers. Enjoy!

10 things you might hear in R&D at MO BIO:

10. I need MRSA tomorrow. Carl, can you start a culture?

9. I brought in those moldy strawberries to test with PowerFood. Have fun squishing that up in the BagMixer^®.

8. I am going to the beach to collect water samples. See you in a few hours.

7. That biofilm from your shower worked great! Can you let it grow a little more next week?

6. MO BIO’s lawn gives the best yields of soil DNA. When in doubt, use the lawn.

5. Pete, can I take some of your blood today for some PAXgene tube stabilized blood RNA preps? You always give me the highest yields.

4. Paraffin Removal Reagent smells great! It’s like aromatherapy for the lab.

3. There’s no smell. I only opened the BME for one second.  Mark, it can’t be coming through the vent in your office unless it’s connected to the fume hood.

2. Craig Cary and Charlie Lee sent some new pictures from Antarctica. Check it out!

1.  R&D meeting is cancelled today. Surf’s up!

***

We have a lot of different projects going simultaneously in R&D,  with products that are getting ready to go to market, some in the research phase of development, and others  just getting started with feasibility studies. And we also collaborate with other researchers who have expertise in areas we don’t and want to be a part of the commercialization of something new.

What makes working for MO BIO so exciting and so much fun is talking to people like you and having the chance to create something that makes life easier for everyone we meet and hearing your feedback in emails and at conferences.

This holiday season, MO BIO is grateful to all of the scientists who inspire us to invent new products and develop new methods. We keep learning, pushing ourselves and having fun because of you.

So we’d like to thank you for a great 2009 and warn you to get ready for an incredible 2010. Some of the new products we have in development are going to be the best we’ve ever produced.

Happy Holidays to All!