About the author

Suzanne Kennedy

Suzanne is Director of R&D at Mo Bio Laboratories in California, and the author of their blog, The Culture Dish. She has a PhD in Microbiology and Immunology from Virginia Commonwealth University.

To enable tagging you will need to register on Bitesize Bio. We're sorry for the inconvenience, but it's free, only takes a few seconds, and it will enable you to view our seminars for free, ask questions from the professional community, and take part in the lively community of Bitesize Bio

We received the following question from Bitesize Bio reader, Beheroze Sattha. It relates to a problem with absolute quantification using plasmids for standard curves. Since many people use this technique it is an interesting one question for us to explore, and it also gives us a great opportunity to cover some important tips for performing qPCR with a new template for standard curves.

Question:

I would like your help to assign copy numbers to plasmid standard dilutions. I cloned a portion of the mitochondrial D-loop gene and after plasmid prep of a single clone made serial dilutions (1:10). In my lab previously they would do a lightcycler run on those serial dilutions using SYBR Green and then assign copy numbers based on the crossing points. The theory used was that a crossing point (CP) of 32.7 would be equal to 10 copies. So the dilution closest to CP of 32.7 would be 10 copies with increasing 10 fold copies for the earlier dilutions (because I had made 1:10 serial dilutions: 1:10, 1:100, 1:1000, etc.

Then I learned from a co-worker that it is better to use the following website:
http://www.uri.edu/research/gsc/resources/cndna.html

So I entered the nanodrop reading of the undiluted plasmid standard and the length of template (TOPO vector and length of my insert) and got the copy number.

The problem is that there is a 10 fold difference between these two methods. 1:1000 dilution of the plasmid standard gives me 8.44E7 copies using the website and 9.99E6 using lightcycler CP.

******

Thanks for your question!

There are a couple things to keep in mind here and I think you’ll be able to solve this problem.

1- The standard curve results and crossing point or Cq numbers are not going to be identical every time. You didn’t mention whether the original standard curve data was performed with the cloned gene or with gDNA or with the PCR product for the gene itself.  If the original standard curves were determined with one type of template and now you have a plasmid template, there will be a difference. Even a change of 1 cycle can have a big effect in absolute quantification.

2- Have you checked the efficiency of the plasmid template? How does it compare to the efficiency of the template used to generate the previous standard curve? If they are different, the quantification will be different.  Ideally you want the efficiency to be above 90%. If the efficiency is below 80%, you won’t want to use this data and you may need to redesign the primers or optimize the chemistry or running times.

2- Every time you order new primers or a new enzyme kit (of a different lot#), you will want to repeat the standard curve results because the numbers can shift. As long as PCR efficiency is high, the data will be accurate, but the Cq may very well be different with new reagents.

3- If you used a plasmid for a standard curve, did you linearize it for qPCR? Many people report that using supercoiled plasmid for standards can cause some variance in results. Try linearizing it first. Here is a recent publication on the subject.

4- When calculating the copy numbers, you may use the length of the PCR amplicon or the entire plasmid. When using the website above, if you use the size of just the amplicon as your input, make sure to adjust the amount of DNA going in to reflect the proportion of the plasmid (see the example in the comments).  If you do not, your reading will be 10 fold off.

5- Make sure your negative controls are negative. Working with plasmids can be tricky because they can easily contaminate solutions. Make sure you have negative controls that are not amplifying because this will boost the real samples and result in inaccurate quantification.

6- Always do the melt curve analysis when using SYBR green and make sure you amplified a single product. Amplification from dimers will add fluorescence and result in an artificially low Cq. You can remedy this by performing an extra data acquisition step at a temperature above where the dimers melt and below where the real product melts. Alternatively, you may want to redesign the primers if dimer formation occurs even in samples with the highest amount of plasmid.

7- With plasmids, it is easy to overload the reaction and have Cq values so early that the detection won’t be accurate. Some instruments have a pre-set baseline setting where they subtract any fluorescent signals generated too early, assuming it is background noise. If you have too much signal in cycles 1-10, this can happen. You don’t want to have samples coming up early so dilute until the first sample has a Cq of 15-18. The subtraction of strong fluorescence in the early cycles will cause all of the data from the more dilute samples to shift right, causing later Cq values than what they are.

Summary:

When using a new plasmid as a standard in qPCR, do an efficiency check  first and compare to the efficiency of the previous assay. For best results, an assay needs to have >90% efficiency, although there are formulas you can use that normalize for differences in efficiency. It is not uncommon that the Ct values are not exactly the same from user to user and from assay to assay for the same gene but with different primers. Just make sure that when calculating copy numbers, you are using the length of the template being amplified and not the entire plasmid which is probably 30 fold bigger than the template and could be the cause of the 10 fold difference in copy number results between the two methods you are comparing. Finally, it’s always good to re-check your standard curve with each new purchase of reagents to make sure no new variables are introduced that could throw off the quantification and make months of work unusable.

About the author

Suzanne Kennedy

Suzanne is Director of R&D at Mo Bio Laboratories in California, and the author of their blog, The Culture Dish. She has a PhD in Microbiology and Immunology from Virginia Commonwealth University.

To enable tagging you will need to register on Bitesize Bio. We're sorry for the inconvenience, but it's free, only takes a few seconds, and it will enable you to view our seminars for free, ask questions from the professional community, and take part in the lively community of Bitesize Bio

We all are very familiar with the effects of cannabinoid receptor stimulation on the body. Relaxation, pain relief, and increased appetite probably come first to mind. These psychoactive effects result from activation of  the CB1 receptor found on cells in the brain by tetrahydrocannabinol (THC).

But there is another receptor, called CB2, that can bind THC and other natural ligands for the cannabinoid receptor. The CB2 receptor is found on cells comprising the immune system and have a multitude of anti-inflammatory and immunosuppressive effects upon activation. In most cases, immunosuppressive effects are undesirable, but sometimes that can be beneficial. Today’s article is an example.

New research from the lab of Dr. Guy Cabral at Virginia Commonwealth University shows that stimulation of the CB2 receptor on macrophages inhibits migration of healthy immune cells towards the HIV Tat protein.  Tat is an essential viral regulatory protein used by HIV to stimulate inflammatory responses and wreak havoc in the body. Tat protein looks suspiciously like some of our own chemokine proteins and can bind to a variety of receptors on immune cells causing activation of cascades that lead to migration of uninfected macrophages towards the HIV infected cells.

The paper by Erinn S. Raborn and Guy A. Cabral, published in the January 2010 issue of the Journal of Pharmacology and Experimental Therapeutics is titled: Cannabinoid Inhibition of Macrophage Migration to the Tat Protein of HIV-1 is Linked to the CB2 Cannabinoid Receptor.  The authors used a macrophage cell line in a migration model system to demonstrate very specifically that when the CB2 receptor is stimulated, macrophages no longer respond to the Tat protein.  The chemoattractant effects are abolished.

Here is an summary of their work.

Introduction:

Macrophages are the primary target for HIV infection and once infected, cells begin producing viral Tat (trans activating factor) protein and GP120 protein in addition to stimulating the production of cellular cytokines and chemokines that induce changes in the immunoregulation of the host. The HIV Tat protein has an additional role of acting as a potent chemoattractant for monocytes, thus contributing to the spread of infected cells.

Most drugs of abuse (opiates, cocaine, amphetamines and cannabinoids) have an adverse effect on immunity, increasing susceptibility to infection. The cannabinoids in particular have been shown to have anti-inflammatory properties, downregulating some of the chemokines and cytokines involved in stimulating macrophage to migrate to infections and inhibiting macrophage function.  Cannabinoids have also been shown to down-regulate the expression of chemokine receptors, notably CCR5, one of the co-receptors used for HIV entry into cells. Thus, a link between the potential anti-HIV effects of the CB2 cannabinoid receptor has been established.

The purpose of this study was to determine whether cannabinoids exert any effect on the chemoattractant properties of Tat in macrophages. In the presence of cannabinoid agonists delta-9- tetrahydrocannabinol and CP55940, human macrophage-like cells (U937 cells) were inhibited from migrating towards Tat protein and this effect was due specifically to CB2. The results show a clear link between CB2 and the ability of macrophages to respond to the HIV protein in a cell culture system.  This work provides the basis for a novel therapeutic target for preventing or reducing HIV associated immunopathology and dissemination in vivo.

Materials and methods:

Cells: The human leukemic monocyte cell line U-937 was used.

Drugs: CB1 and CB2 receptor agonists used were: THC and CP55940. The CB2 specific agonist was O-2137-2 and the CB1 specific agonist was ACEA. The CB1 and CB2 receptor antagonists were SR141716 (SR1) and SR144528 (SR2), respectively. The full names of the drugs and the Ki information is described in the paper.

Tat: Recombinant human HIV Tat protein was obtained from Immunodiagnostics, Inc.

Cell Migration Assay: 35 mm tissue culture plates with upper and lower compartments separated by a polycarbonate 8 micron pore membrane were used. Drug treated  or control treated U937 cells were incubated on the top chamber and Tat protein or serum-free media plus vehicle was in the bottom chamber. Migration of cells to the bottom chamber was visualized with an Olympus CK2 inverted microscope connected to a digital video camera. The number of cells were manually enumerated. A greater than 2-fold increase in the number of cells in the presence of chemoattractant compared to no Tat was a positive response. The EC 50 or inhibitory concentration was the concentration of cannabinoid that results in a 50% reduction in macrophage migration.

Knockdown of CB2 expression using siRNA and RT-qPCR:   siRNA for the CB2 receptor was used for transient transfection of cells and knock-down shown using Western blot analysis.  SYBR Green was used for qPCR analysis after reverse transcription of RNA, qPCR was performed using the SmartCycler. Full details of the experimental design for qPCR and transfection are provided in the paper.

Results:

To begin, the authors first confirmed the expression of the CB2 receptor in U937 cells, both on the RNA and protein level. The CB1 receptor, typically expressed in cells of the brain, was not found in U937 cells using RT-qPCR. Their second set of confirmatory experiments proved that the U937 cells would migrate in response to Tat protein as has been previously described for human monocytes in blood.  The migratory response was maximum at 50 nM Tat and so the authors used this concentration for their studies.

The next experiments looked at the  migration of macrophages in the presence of the different drugs. All results were compared to vehicle controls (ethanol was used to dilute the drugs). Using vehicle alone with Tat protein in the bottom compartment, migration was the same as with no vehicle. When cells were treated with THC, however, migration of macrophage was inhibited by 50%. And using the agonist CP55940, migration was inhibited by 58%. Using the CB2 receptor specific agonist, the same effect of >50% inhibition was observed, however, but not suprisingly, using a CB1 receptor agonist, ACEA, had no effect on migration.

These results clearly indicate that the CB2 receptor on the human macrophage-like U937 cells plays a role in migration in the presence of the HIV Tat protein and this effect can be blocked when the receptor is activated.

To confirm the effect of migration was due to the CB2 receptor, the receptor antagonists were used to demonstrate that when signalling through the receptor is blocked, the effect is reversed.  The antagonist will bind the receptor but not activate the signalling cascade in the cell, thus blocking it from being activated by the ligand CP55940. Using the CB2 specific antagonist SR2 alone, migration in the presence of Tat is the same as controls- it is not inhibited. When CP55940 is combined with the SR2 compound, inhibition of migration is reversed.  SR2 prevents the protective anti-migration effect of CP55940.

To further prove the role of CB2 in this effect, siRNA mediated CB2 knockout cells were employed in the cell migration assay. The authors confirmed that neither the transfection reagent nor the siRNA itself had any effect on cell migration. Using the CB2 knockout cells in the presence of THC, migration was observed similarly to untreated cells. This is consistent with the results of blocking the CB2 receptor with antagonist. The CB2 receptor was not available for activation by the THC and thus no protective anti-migration effect was observed.

Discussion:

Chemokines are cytokines that function by directing the flow of inflammatory cells to sites of injury or infection in the body.  HIV uses our immune system cascade meant to protect us for its own benefit, by producing the viral Tat protein to bind the receptors meant for chemokines and stimulate the migration of more healthy macrophages to the HIV infected cells, increasing their opportunity to spread. The stimulation of chemokines also increases the number of receptors on the surface, making it easier for HIV to infect new cells.

The ability to slow down the migration of healthy cells towards HIV infected cells and to down regulate the expression of chemokine receptors would slow the progression of HIV associated disease. In this paper, the authors demonstrate a connection between the cannabinoid receptor on immune cells, CB2, and the migration of macrophages towards Hiv Tat protein. When the CB2 receptor is stimulated with an agonist, migration is inhibited. Whether this is due to down regulation of the chemokine receptors used for binding or due to down regulation of chemokines from CB2 activated cells has yet to be determined.

But the fact remains that the cannabinoid receptor CB2 may be a therapeutic target for preventing hyperactivation of the immune system by HIV and potentially, in the future, to help stop widespread infection.

Final note:

I found this paper to be an exciting advance in the fields of HIV and drugs of abuse. A therapy involving cannabinoids would be far less toxic than current drugs used to stop the progression of HIV. With all the ways HIV has found to use our own immune system against us to survive, an approach that counteracts HIV by taking back control of the immune system could be far more effective than developing toxic drugs that target HIV directly or using cytokines that further activate a tired immune system.

About the author

Suzanne Kennedy

Suzanne is Director of R&D at Mo Bio Laboratories in California, and the author of their blog, The Culture Dish. She has a PhD in Microbiology and Immunology from Virginia Commonwealth University.

To enable tagging you will need to register on Bitesize Bio. We're sorry for the inconvenience, but it's free, only takes a few seconds, and it will enable you to view our seminars for free, ask questions from the professional community, and take part in the lively community of Bitesize Bio

This past week I found myself asking this question quite a few times. What is going on with the peer review process? Is anyone actually reviewing the papers getting into journals anymore?

This is due to some recent experiences I’ve had with papers published in both the larger highly reputable journals and smaller niche journals that has left me wondering.  I review papers for Current Issues in Molecular Biology and I have run the peer review process for them so I understand what is expected as an editor or a reviewer. The journal is counting on me to ensure that a high quality paper is approved and nothing less. As an editor running the peer review process and selecting reviewers of the paper, it is my responsibility to make sure I choose people who will be fair, unbiased, and have the time to actually read the paper.

I am incredibly busy, but this is a responsibility I take seriously because not only does a poorly written paper reflect badly on the journal and on the authors, it reflects on the editor.

How did this get published?

Recently I came across a paper that was published in a popular journal for microbiologists despite the fact that it had no control experiments, the conclusions didn’t match the data (see the previous article on the importance of reading a paper thoroughly), and the entire study missed the opportunity to make any scientific contribution to the field by focusing on the wrong points.  How did this make it through peer-review, I asked myself?

Letters can be written to the authors and the editor, however, it doesn’t correct the problem. How does a scientific study gone awry get published anyway? How does every reviewer miss such obvious flaws in a paper?

More examples…

Recently I read a paper from a smaller niche journal for microbiologists that a scientist/friend asked me to comment on. We both came to the same conclusions: the paper was garbage. The authors used the incorrect name for their organism, making one up that doesn’t exist and mislabeled a figure so that the results section did not match the figure legend, making interpretation confusing. Furthermore, they left a key piece of information out of their methods (how much bacteria they used to inoculate the soil), so the data obtained was uninterpretable, and to cap it all they misinterpreted their very own data in the conclusions.

How does this get published? Was it reviewed at all? A paper of this low quality brings down the journal and its editors, and affects the whole field.

It’s not just microbiology journals!

The popular science and general methods type journals are not immune to this problem. Several months ago I read an article in a very popular magazine that used qPCR for their study and attempted to use the MIQE guidelines to validate their work. It was great to see people attempting to use the guidelines.

However, upon opening the attached spreadsheet in the Supplemental Data, I was shocked to see that most of it was empty. The authors actually wrote “NA” in almost every field of the MIQE checklist and didn’t provide any information on yields, purities, etc.  How many people do you think took the time to go to the Supplemental Data and actually see the checklist? I’ll hazard a guess that it is very few since no one else noticed the alarming lack of information contained in their “supplemental data”.

I know we are all busy and I know that we want to help our colleagues to get published, but there needs to be more measures in place that ensure that the standards are high. 

Here’s what I suggest

My suggestion is that if the journal allows the authors to choose reviewers for the paper, the editor running the peer-review process should use only one name from their list and the others are not. Chances are that the reviewers suggested by the authors are going to be very lenient (after all, they are going to ask for the return favor when they publish their next paper) and the one with nothing to gain by approving the paper will give a more accurate assessment.

And editors need to stay objective to the body of work regardless if the authors are friends or collaborators.  It is just like referring someone for a job- it reflects on us as editors if we recommend a piece of work that really is not up to par. No journal wants to publish papers that need erratum published later, or worse, no one takes the time to let the journal or authors know that an article needs erratum and instead the community decides that that the journal is too low tier to even publish there or that the people who publish there are those who get rejected everywhere else.  And of course, no one wants to cite a paper with obvious flaws so the citation index continues to go down.

Ok, so maybe it’s not ALL broken…

I know that the peer-review process does work successfully and that there are many journals making sure that their review process is stringent. It’s probably no coincidence that these journals usually have the highest citation indexes as well.

My advice to all the readers out there preparing papers for submission to journals is to step back and look at your data as objectively as you can. I know you have a theory or model you are trying to prove and want the data to support you. So knowing that, step back and read it again from an outside point of view. What other interpretations could there be? What else could be going on? Let the data tell you what is going on.  Be open to other interpretations of the data and write it up objectively.

Not only will you have a stronger paper that more people will cite, but, it will mean that you were the first to think of it and when other people follow your lead and build on your work, they will have to say you suggested it first. 

And don’t be disappointed if you need to make revisions to the paper when it comes back for review. Thank the reviewers for taking the time to read your work carefully and know that you will be able to re-submit a much stronger and more citable paper in the next round.

About the author

Suzanne Kennedy

Suzanne is Director of R&D at Mo Bio Laboratories in California, and the author of their blog, The Culture Dish. She has a PhD in Microbiology and Immunology from Virginia Commonwealth University.

To enable tagging you will need to register on Bitesize Bio. We're sorry for the inconvenience, but it's free, only takes a few seconds, and it will enable you to view our seminars for free, ask questions from the professional community, and take part in the lively community of Bitesize Bio

Recently we received a question from Bitesize Bio reader Sonia after our article How to: Get Better Plasmid Midiprep Yields. She asked: “What could be the problem when one sample gives a good yield while the other plasmid gives poor a one, when both the samples were processed simultaneously, and in the same way.”

This is a good question because many things can cause differences in yields between plasmid preps.  Let’s resolve this mystery one point at a time and go over some reasons why you might get low yields when you prep plasmids.

1. Plasmid backbone

You prepped two plasmids simultaneously using the exact same protocol and had different yields. If the plasmids have the same backbone (that is, they are both pUC, or pBluescript, etc.) then the reason leans towards the insert playing a role. Some inserts can be problematic for bacteria. It might be that a protein is made that makes the bacteria sick (for example, DNase) or it could be that the insert is unstable (for example, repetitive sequences). To overcome the problem, try using a specialized competent cell line. For unstable inserts, try the STBL2 cells from Life Technologies and for growing clones with toxic proteins, try the T7 Express LysY/Iq Competent cells from NEB.

Another important point is how the insert size changes the copy number of the plasmid. Large inserts will reduce the number of copies of the plasmid.

2. Copy number

If the genes are cloned into different vectors then the issue could be that the plasmids are replicating at different rates. One may be high copy and one may be a medium or even low copy plasmid. Some examples of  low copy plasmids are ones using the backbone pBR322 and pACYC, which are older and not used often in cloning work today. Many vectors used for protein expression are medium copy. This is desireable because when producing proteins, sometimes if growth is too fast, it enhances the chance of the protein becoming insoluble or forming inclusion bodies.

3.  Culture issues and antibiotic

Nick went over the technique for growing bacteria to obtain a healthy culture in late log phase where you can get the most plasmid in our original midiprep how-to article.  This technique works well as does using a 1:1000 dilution of starter culture into a large scale culture (so 100 ul into 100 ml) if you want to grow the bacteria overnight for 12-16 hrs.  One important consideration is the antibiotic. The bacteria are going to break down the antibiotic while they are growing in the culture. If not enough antibiotic is added or if the stock is old and not at the correct strength, the antibiotic selection pressure may not last very long and you could end up having a culture that was antibiotic-less for most of the culture time. Plasmid yields will go down without the selective pressure to keep it.

As a reminder, the state of the culture is critical for high yields of plasmid. For maximal yields, the culture should be in late log or early stationary phase. If the culture overgrows, you will be harvesting more dead bacteria than live cells and this also leads to genomic DNA contamination in the prep. If the culture is undergrown, then of course, yields are lower than expected. You can mistakenly undergrow a culture by using old colonies from plates or starting direct from a frozen stock and not from a colony. The lag time for the bacteria to ramp up is much longer when you use either of these approaches.

Streak fresh colonies:

One more point that people forget when setting up their starter cultures or overnight cultures is the age of the plate you are using to pick your colony. If your plate is old, you may have picked a nice big colony but it will not be all living cells. And if there were satellite colonies sitting around the original colony where the antibiotic no longer exists, those will not have plasmid and will be introduced into your culture. So streak a fresh plate before starting to ensure the best result.

So let’s assume that Sonia’s plasmids were the same vector, same antibiotic, grown exactly the same, and each have different inserts that are not too large or unstable or toxic. What else can it be?

4. Processing steps:

If the culture conditions are not the problem, then we have to look at something with the downstream steps. Since we are talking about Midipreps, let’s discuss what can go wrong with the anion-exchange procedure next.

Nick covered alkaline lysis in great detail and typically these reagents in the kits are stable and fine. Solution 2 (the one containing NaOH and SDS) can break down over time with exposure to air, but in general, they work for lysing bacteria for the life of the kit.

The other area of plasmid preparation where DNA can be lost are the final steps after anion-exchange which is the final precipitation step in isopropanol and finding the DNA pellet.

Isopropanol quality:

Many labs have isopropanol in large containers that have been opened and closed over the course of a year. For the best result in the precipitation step, make sure the isopropanol used is not the old bottom-of-the-barrel stuff. Use some isopropanol from a new bottle or a smaller bottle that is not who-knows how many years old. This makes a huge difference in the size of the DNA pellet you obtain after centrifugation.

Don’t lose the pellet!

Isopropanol pellets are glassy and clear and difficult to see. The best practice is to mark the side of the tube where you expect the pellet to form after centrifugation in a fixed angle rotor so when you decant the isopropanol, you know where to look for it. Keep an eye on the spot and look for the glassy material. Sometimes this is difficult because many people use the oakridge plastic tubes which are opaque. If you have glass corex tubes, this is a nice alternative and they can be baked to make them pyrogen free.

Sometimes, if you have concerns about losing the pellet, it is good practice to pour the isopropanol supernatant into a 15 ml tube to save it, just in case the pellet slipped off the wall. But this does not normally happen as long as you do not let the sample sit for long after the centrifuge stops. Once it is done, be right there to decant the sample. The only times I have seen a pellet come loose from the wall is when I was late getting to the centrifuge and it sat still for a few minutes.

Whether you use Oakridge tubes or glass, just note where that pellet should be. Once you wash with 70% ethanol, the pellet becomes visible.  When you are ready to resuspend your pellet, you’ll know exactly where to find it because you marked the tube.

Caution! It is not always a pellet!

Sometimes with fixed angle rotors, the DNA may not always form a nice tight pellet at the side wall. It can sometimes smear down the side. For this reason, I always use my resuspension buffer to wash down the side of the wall above my pellet to make sure I solubilize every molecule of plasmid that may be present even though I can’t see it.

It is a shame to do all that work and then lose the DNA pellet right at the end! More tips on perfect DNA pellet recovery is found here too.

We had a nice discussion about DNA precipitation in a previous article and it was the consensus that the most important factor in obtaining high yields is centrifugation speed and time. Don’t cut the centrifuge time short unless you can turn up the speed.

I thought it would be good to mention that many plasmid kit manufacturers have recognized that the pelleting step is problematic for some users so have developed kits that desalt the DNA using “precipitators” or silica disc filters. These are a fast alternative to centrifugation. However, you still need good isopropanol for these to work so always use fresh.

Summary:

Large scale plasmid DNA preps have a lot of steps where things can go wrong but in my experience, the problem is usually either the culture or the DNA precipitation.  To check if your culture is healthy, just take 1-2 mls out of your 100 ml flask and then do a quick miniprep on it to see how much plasmid/ml is there. That will give you a good idea of what you will get from the rest of the sample.

So remember to get great plasmid yields, do a little background first on your vector and insert to make sure there is no reason for the DNA itself to be a problem and then start with a fresh colony and a starter culture and fresh antibiotic. And at the end, fresh isopropanol will be key to a thorough precipitation of all the plasmid DNA.

That about covers it folks. If you have any more questions, problems, or concerns, send them our way and we will put our heads together to figure out how to help!

About the author

Suzanne Kennedy

Suzanne is Director of R&D at Mo Bio Laboratories in California, and the author of their blog, The Culture Dish. She has a PhD in Microbiology and Immunology from Virginia Commonwealth University.

To enable tagging you will need to register on Bitesize Bio. We're sorry for the inconvenience, but it's free, only takes a few seconds, and it will enable you to view our seminars for free, ask questions from the professional community, and take part in the lively community of Bitesize Bio

Who is Andy Bass?

Answer: A surfing scientist turned businessman from Alabama with a passion for educating the world about molecular biology.

Andy is the CEO and founder of Yonder Biology, a brand new biotech company located in northern San Diego county here in California. I had a chance to speak with Andy about his new venture into personalized DNA art. I was really impressed with Andy’s passion for bringing science to the public and using his new company as a platform to further educate people on how science is impacting their lives.

You may recall Alex’s pre-holiday’s Bitesize Bio article on dealing with family stress when you’ve had to explain over and over “I’m not that kind of doctor!“.  Yonder Biology aims to help in bridging the gap between the kind of doctor that does molecular biology with the rest of the world and for him, the bridge begins with molecular art.

Andy received his bachelor’s degree from the University of Alabama, Birmingham and went to work in biotech sales for Life Technologies, back in the pre-Invitrogen days.  Ten years after graduating college, he is ready to embark on his own frontier, taking a leading role in developing ways to make science understandable for non-scientists.

In my interview with Andy, we discussed his plans for the new company and advice he has for budding entrepreneurs who think they have a cool idea and want to try to be their own boss.

Tell us about your background?

On the science side I have a bachelors of science in molecular biology from UAB and have worked in the lab in industry as a associate scientist. On the business side, I worked in business development and sales. I’ve always been entrepreneurial and creative. While that hasn’t been my career, it is always my driving force. Now with Yonder Biology I get to blend the mix of creativity and science.

What were you doing before you started this business?

I was working for Bio-Rad in sales and I covered southern California and all of the pharmaceutical and large biotech accounts. Before that I was working for Sigma Aldrich in sales.

What gave you this idea to make art out of DNA?

I called a couple friends who have the creative and entrepreneurial drive too and when you explain to people not in science what you do, it’s always interesting and they have a hard time grasping it. We thought about how to bridge the cutting edge research going in in life science industry with the every day consumer: “mom and dad”. DNA art is a simple but meaningful way to involve them in science.

I really wanted to blend creativity and science together into a product and looking at what is out there now, we saw personal genomics, DNA chips, and next generation sequencing as areas where we wanted to be involved. But we took a step back and said “OK what can we do right now?” So what we did was create the personalized DNA art portrait and this is just the beginning. We are working towards incorporating next gen sequencing into our offerings and marketing that to the everyday consumer.

And not necessarily in art. We have bigger dreams. We are starting on the creative end but want to bring science to the average person.

Looking forward, we see this as a small educational step, where mom and dad can see what DNA looks like, to when personal genomics really enters the mainstream market and doctors are making decisions based on your genome. One area we want to go into is finding ways of presenting life science research to the public. So an education role is one option.

How did you get your company started?

We got a core group of people together who were all interested in these same goals and invested our own money as well as put some ads on Craigs list to get the core equipment, like a thermocycler and basic gel equipment and we built our own imaging system. We received a small amount of angel investor money to help us out. Currently we have a lab only and are looking at a few spaces where we can merge a lab with something like an art gallery.

How hard was it to get started? What does it take to start your own biotech company?

We got it up and running pretty quickly and we thought it was as easy as run PCR, run a gel, take a picture. Then we realized we had to design a website that was search engine optimized, we had to come up with a way to collect the DNA, then how we will package the final product, etc. So there are all of these little things that you need but don’t necessarily anticipate or think about when you first start out but you need answers for.

I would say that you don’t know the questions until you get started, when it is your first venture.

Where does the name Yonder Biology come from?

Interesting story.  I was out on my surf board here in Carlsbad sitting in the water with friends I grew up with from Alabama and we were pointing to something in the water and I said it was ”over yonder” and then I said it again. I went back and proposed it to a couple of the guys. I asked what  “yonder” means to them and they said “the horizon”, “beyond”, and “the future”.  We really liked these three themes on what “yonder” meant to us. So we stuck Biology on the end and we had our name.

How does it work? What methods are you using in the lab?

It is DNA fingerprinting. We’ll send out a DNA collection swab and you’ll rub on the inside of your cheek, send us back the swab in an envelope, and we’ll extract the DNA. We test the purity (260/280 ratios) and then we’ll do PCR to amplify the STRs. The specific primers pairs we use we are not disclosing but will tell people the general chromosome locations. We use particular ones and orient them the same on all the gels.

After gel electrophoresis and staining, we take the picture with our custom imaging system. The unique thing about our imaging system, is that it is so high resolution, we can essentially put the picture up on the side of a building without losing any quality.

We have experimented between saliva samples and swabs and we get a lot more DNA with saliva but we weren’t sure what we would get back from people so we thought it was more sanitary to use a swab. The DNA yields are more than enough.

Did you have any qualms about moving from hard science to science art?

Our company hasn’t taken our eye off of hard science. We see that as the core of what we are doing. For example, if any issues come up in the lab, we have to be scientists and use our background to solve those problems. And looking forward we want to grow in the personal genomics space so we are up on cutting edge technology such as whole genome amplification and next-gen sequencing technologies. While what we do is for consumers, we plan to make our name in personal genomics in the future.

How did your family and friends respond to your plans to quit your job and begin this venture?

Everyone that knows me probably thinks I am a little bit crazy anyway? But everyone in my family knows we have a plan for the company and we have milestones to make sure we are moving in the right direction so it’s a very calculated risk and my friends and family have been very supportive. We set timelines in place where we want to have certain things accomplished and if we are not on track, what do we need to do.

Have you had any unusual or fun requests (ex. pets)?

We recently opened up our DNA art to pets so we’ve done a handful of dog DNA portraits. We’ve also had a request for whole genome sequencing for whales. One other thing we’ve done is launch “Duet” and “Quad” DNA portraits- so a couples portrait or family portraits. Currently we can combine up to four profiles on the same canvas.

What advice do you have for other entrepreneurial scientists who think they have a novel idea and want to start a biotech company?

Surround yourself with the right people. You don’t have to know everything but it’s good to have people who know the things you don’t. Diversity is good. Going from web design to PCR, to even the legal aspects its good to have people who can contribute different aspects.

Also things sometimes take longer than you expect or cost more than you expect so you have to be prepared. My advice is to take a conservative approach when looking at financials and sales projections.

Thanks Andy!

It sounds like Yonder Biology has clear direction and goals and an exciting future ahead. At the dawn of personalized medicine, a company like Yonder Biology will be at the forefront of educating the public.

If you live or work around the San Diego area, Andy will be exhibiting at the San Diego Science Festival March 27th and will have some DNA portraits of UCSD professors on display at The Loft.

You can follow Andy on Twitter at http://twitter.com/yonderbiology

Let us know what you think about DNA art. And if you have any questions for Andy about his products, or on starting a biotech company, please leave us a comment.

About the author

Suzanne Kennedy

Suzanne is Director of R&D at Mo Bio Laboratories in California, and the author of their blog, The Culture Dish. She has a PhD in Microbiology and Immunology from Virginia Commonwealth University.

To enable tagging you will need to register on Bitesize Bio. We're sorry for the inconvenience, but it's free, only takes a few seconds, and it will enable you to view our seminars for free, ask questions from the professional community, and take part in the lively community of Bitesize Bio

When I worked in technical service for a well known biotech company, I have to confess that we often used a certain phrase in the frustration of dealing with calls from angry scientists ranting about a problem they were having with a kit because, as it turned out, they didn’t read the manual.

“Read the F***ing manual” (RTFM), was the phrase (only used after we put the phone down of course). A bit naughty, but it was certainly an essential stress reliever! This article today is not about reading manuals, but research papers.

It is my appeal to everyone to RTFP.

I find that there is a growing problem, especially amongst newer members of the scientific community, of people reading ahead to the conclusions of the paper and taking them as fact without having read the methods and results  sections, or critically analyzing the data.   This is very poor practice for many reasons but the main point is that just because the article was accepted by the journal you should not assume that the work was reviewed stringently, carried out correctly or reported objectively.

The conclusions contain the take-home message of the paper but these other sections are just as, if not more, important. Here’s what you should look out for in each section.

Introduction:

The introduction builds the story and explains what previous research has shown and what this new research will add to the current knowledge base. This section helps you to determine if the authors did a thorough review of the field, and if it’s your field, you (should) know whether the authors left out any particular papers that are important to cite. If key papers are left out of the introduction, how careful were they with the rest of the paper?

Methods section:

The methods section should clearly and thoroughly outline exactly what was done.  Read it carefully. Are the controls described? Did they modify commercial kits, and if so do they explain how? Are they doing the right comparisons? Did they include enough data points?

If the data is qPCR, then take the time to look even more carefully. According to the MIQE guidelines, the authors need to explain the nucleic acid purification method, yields, and purities, which kits they used, how they determined the efficiency of their assays, and how many replicates they did. There are a lot of factors that can influence qPCR data and if the paper is leaving out some of the information, you can’t make accurate conclusions on the data.

Results section:

Here is the part where the authors interpret their data. Each figure is reviewed one by one. Read this part critically.  How do the controls look? How do the qPCR curves look? Are the Westerns clean? Is all the data in graphs and tables instead of allowing you to see how it actually looks? You do these experiments too and you know how data should look.  The quality of the data is as important as what experiments they did.

Conclusions/Summary:

Here is where the authors have the chance to pontificate on their work and tell you what they think it means.  They are making their conclusions based on the results. Now if you have read the whole paper, you are in a position to either agree or disagree. Do you agree with how they interpreted the data? Can you think of alternative explanations for their results? Are they being objective? You’ve looked at the results and you’ve reviewed their methods.  What do you think?

As scientists we all have our theories and we want our data to fit our model.  We want to be right. Sometimes the need to be right overrides accuracy.  It is human nature. I once had a PI tell me “If you want to prove me wrong, go find another lab”. The data didn’t fit his model and he wasn’t open to changing it, which was bad news.

The message in this article today is to please read your papers. Please, please do not just read the conclusions and take them as truth. They aren’t always the only explanation – you may not actually agree. Besides critical review of scientific papers is a necessary skill and will serve you well. Not only as a future reviewer of journal articles, but as writer of your own research.

Let us know in the comments — do you RTFP?

About the author

Suzanne Kennedy

Suzanne is Director of R&D at Mo Bio Laboratories in California, and the author of their blog, The Culture Dish. She has a PhD in Microbiology and Immunology from Virginia Commonwealth University.

To enable tagging you will need to register on Bitesize Bio. We're sorry for the inconvenience, but it's free, only takes a few seconds, and it will enable you to view our seminars for free, ask questions from the professional community, and take part in the lively community of Bitesize Bio

Happy New Year!

Now that 2010 has begun, it’s a good time to start thinking about how you want to spend your travel money (or which meetings you want to nudge your PI about spending their travel money on for you). So to help you plan, here is a list of the bigger, more popular conferences in the life sciences in 2010.

These conferences all take place in the US so please feel free to send us your comments and let us know what additional conferences, large and small, you’ll be attending in the rest of the world that you think are providing the latest and greatest information for your field of work.

January

Plant and Animal Genome (PAG)
January 9^th-13^th
Town and Country Hotel, San Diego CA

LabAutomation 2009
January 23^rd-27^th
Palm Springs Convention Center
Palm Springs, CA

February

American Academy of Forensic Sciences 62nd Annual Scientific Meeting
February 22^nd-27^th, 2010
Washington State Convention & Trade Center
800 Convention Place
Seattle, WA 98101

Biophysical Society’s 54th Annual Meeting
February 20^th-24^th
San Francisco, CA

March

American College of Medical Genetics Annual Meeting
March 24^th-28^th
Albuquerque Convention Center, Albuquerque, New Mexico

April

American Association for Cancer Research 101^st Annual Meeting
April 17 ^th-21^st
Walter E. Washington Convention Center, Washington, DC

Experimental Bio 2010 Annual Meeting
April 24^th – 28^th
Anaheim Convention Center, Anaheim, CA

May

American Society of Microbiology, 110^th Annual Meeting
May 23^rd-27^th
San Diego Convention Center, San Diego, CA

July

Plant Biology 2010
July 31^st- August 4^th
Montreal, CA

American Society of Virology 29th Annual Meeting
July 17^th-21^st
Montana State University, Bozeman, MT

November

Society for Neuroscience 40th Annual Meeting
November 13^th -17^th
San Diego Convention Center, San Diego, CA

American Society for Human Genetics 60th Annual Meeting
November 2^nd-6^th
Washington DC

Association for Molecular Pathology 2009 Annual Meeting
November 17^th -20^th
San Jose McEnery Convention Center
San Jose, CA

December

American Society for Cell Biology 50th Annual Meeting
December 11^th – 15^th
Philadelphia, PA

An alternative to these large diverse conferences are small focused meetings where you will be able to spend more time talking to your field experts and perhaps even share a meal with a PI whose work you admire.  A good place to look for small but focused conferences is the Cold Spring Harbor Labs, the Gordon Research Conferences, and the Keystone Research Conferences. Any of these will give you an intense learning experience.

No matter what you decide, just make sure to go to a conference if you can, even if it’s a local one. Nothing else can invigorate your mind and rejuvenate your passion for your work like a conference.

As well as telling us where you’ll be going in 2010, you could also let us know what was your favorite conference from 2009 and why.

About the author

Suzanne Kennedy

Suzanne is Director of R&D at Mo Bio Laboratories in California, and the author of their blog, The Culture Dish. She has a PhD in Microbiology and Immunology from Virginia Commonwealth University.

To enable tagging you will need to register on Bitesize Bio. We're sorry for the inconvenience, but it's free, only takes a few seconds, and it will enable you to view our seminars for free, ask questions from the professional community, and take part in the lively community of Bitesize Bio

Perhaps at no other time of year like the winter solstice is the mixture of religious beliefs and daily life more intertwined.  Most people, regardless of race and country of origin, come from a faith that believes in God or a Higher Power.

As scientists, it is a widely held belief that we do not believe in God because of our passion for truth. Some think that science and God do not mix and cannot mix; that you cannot be a scientist and actually believe in a higher power or a universal source of knowledge that is not measurable by any lab test.

My experience of what scientists believe

When I speak to my colleagues and friends in the science community, I actually find the opposite belief to be true. Most people I speak to not only believe in God, but in the paranormal, spiritual, and supernatural. We can’t measure any of this with a DNA or RNA test, yet for some, proof is not always something you can hold in your hand. What I have found is that for most scientists, their own experience is proof enough.

This makes sense to me because as scientists, we are trained to have an open mind and to not let our personal biases sway our results.  Scientists need to be open to any and all possibility in order to make progress. So a scientist who has experienced divine guidance or intervention, while knowing that there is no explanation with physical laws on how such a thing could have happened, has all the evidence they need to know fact from fiction.

The stats

The stats say that the split is about 50-50 of those who believe in God and those who do not. A survey taken by the American Association for the Advancement of Science (AAAS) in May and June of this year and reported by David Masci in the Los Angeles Times, found that 51% do believe in God and 41% do not.  These numbers haven’t changed much over the last 100 years either,  despite the numerous discoveries in evolution and biochemistry over the years.

The same poll found that 41% of chemists believe in God while scientists in the fields of biology and medicine were much less likely to believe in God (32%). In terms of age, the younger generation of scientists (18-34) are more likely to believe in God than their senior colleagues.

In comparison, the scientific community tends to believe less that the general public do.  95% of American adults say they believe in a God or higher power and only 32% believe in evolution whereas 87% of scientists believe that life evolved over time.

Based on this data, it would suggest that for many of us, science and religion are not necessarily incompatible and one does not need to choose between the two.  We can believe in a higher power and evolution. We can experience things not explainable by science and not need to write it down in a lab notebook as proof it happened.

Many of our scientific predecessors believed in God; Sir Francis Bacon (1561-1627), Rene Descartes (1596-1650), Robert Boyle (1791-1867), Gregor Mendel (1822-1884), and Albert Einstein (1879-1955). Even under persecution for teaching that the sun was the center of the universe, Galileo Galilei (1564-1642) maintained his faith in God. And Galileo had no proof that his theory was true using the tools available to him at the time.

Brilliant minds over the ages have recognized that there are some things we can’t explain but that doesn’t mean they aren’t true. Sometimes all you have to base your theories on is a hunch or a feeling or indirect evidence that there is more to this than meets the eye.  Does it mean we reject everything that doesn’t fit our cozy model because we can’t measure it in a test tube? No. It means we keep an open mind until we can.

Here’s what I think

I love science because I love the process of solving the riddle and uncovering the clues to unraveling whatever problem I am trying to solve. I love the process of discovery and not just the end result. Fortunately for us scientists, there is an infinite number of puzzles to tackle in the universe and things to discover.  Thank God for keeping a few things secret.

ps: I wanted to let you all know about www.bethematch.org. Your bone marrow may hold the cure for a child with cancer. Joining the registry is easy. It is just a cheek swab.  You don’t need to give bone marrow unless you are determined the best match for a patient.  The greatest present to give any child or parent of a sick child is the chance to live. You might be the match!

About the author

Suzanne Kennedy

Suzanne is Director of R&D at Mo Bio Laboratories in California, and the author of their blog, The Culture Dish. She has a PhD in Microbiology and Immunology from Virginia Commonwealth University.

To enable tagging you will need to register on Bitesize Bio. We're sorry for the inconvenience, but it's free, only takes a few seconds, and it will enable you to view our seminars for free, ask questions from the professional community, and take part in the lively community of Bitesize Bio

Since our most popular article of all time (“The Basics: How Ethanol Precipitation of DNA and RNA Works”) was published, many of our readers have asked us to further explain the difference between precipitating DNA with ethanol vs. isopropanol and which is the better choice. So today, I’ll meet the challenge and discuss the pros and cons of ethanol vs. isopropanol.

First, let’s review what we know about what is needed for precipitation of DNA or RNA with ethanol:

1. Salt to neutralize the charge on the nucleic acid backbone, causing the DNA to become less hydrophilic and fall out of solution.

2. Ice to chill the sample. Lower temperatures promote the flocculation of the nucleic acids so they form a larger complex that readily pellets under the centrifugal forces of a microcentrifuge.

3.  A nucleic acid concentration high enough to force the DNA out of solution (if the conc is not high enough, you can add a carrier nucleic acid or glycogen to enhance the recovery).

4. Centrifugation to pellet the sample

The difference between isopropanol and ethanol is the solubility of DNA in each solvent.

DNA is less soluble in isopropanol so it will fall out of solution faster and at a lower concentration, but the downside is that the salt will too. With ethanol, the DNA needs to be at a higher concentration to flocculate but the salt tends to stay soluble, even at cold temperatures.

DNA falls out of solution in 35% isopropanol and 0.5M salt. Using ethanol, the final concentration needs to be around 75% with 0.5M salt. So for the typical precipitation protocol, isopropanol is added from between 0.7-1 volumes of sample and ethanol is added at 2-2.5 volumes of sample.

The choice of which solvent to use depends largely on the volume of sample you need to precipitate.

If you are precipitating small volumes of DNA, and you can fit the required amount of solvent into the sample tube, then ice cold ethanol is the preferred choice. You can chill it (some people use liquid nitrogen or -80C to accelerate the precipitation) and precipitate more DNA without the salt contamination that would occur from chilling isopropanol. Afterwards you need to wash the pellet with 70% ethanol to remove salt.

Isopropanol use useful for precipitations where you have a large sample volume  (e.g. the eluate you get after using a Qiagen plasmid Maxi Kit) because less solvent is needed, so you can fit the whole lot in the (15ml) tube. But because salts are generally less soluble in isopropanol than in ethanol, they have more of a tendancy to co-precipitate with the DNA. So to lessen the chances of salt precipitation, isopropanol precipitations are carried our at room temperature with minimal incubation times.   Once the DNA or RNA pellet is recovered from the isopropanol, you’ll want to wash it with cold 70% ethanol to remove excess salt and to exchange the isopropanol with the more volatile ethanol. It is ok to chill the isopropanol precipitated sample, if you are sure that it is not excessively salty.

Because DNA is less soluble in isopropanol, isopropanol allows precipitation of larger species and lower concentrations of nucleic acids than ethanol, especially if you incubate it cold and long. If you do this, just remember to wash the pellet several times in  70% ethanol after pelleting, to reduce the amount of salt you carry over.

So how do you choose when to use isopropanol and when to use ethanol?

Use ethanol if:

1. You have room to fit two volumes of ethanol to sample in your tube
2. The sample needs to be stored for a long period of time and will be chilled
3. You need to precipitate very small pieces of DNA or you have a very low concentration of sample so you want to chill it longer and colder.

Use Isopropanol if:

1. You have limited in space in your tube and can fit only 1 volume of sample
2. You need large molecular weight species because incubation at room temperature for short periods of time will not be conducive to precipitating small species of nucleic acid
3. You are in a hurry and want to accelerate the precipitation of nucleic acids at room temperature

What do I prefer? I use ethanol over isopropanol for most cases, but will use isopropanol if I need to make everything fit in one tube. My preferred protocol is 2 volumes of ethanol and freeze at -20C for at least an hour or overnight for best results. I centrifuge the sample at full speed for 20 minutes to make sure I get everything down. I always wash with 70% ethanol and then centrifuge for 10-15 minutes and keep my eye on the pellet when I decant everything. You need to note or mark the side of the tube where the pellet is expected to be and don’t let it out of your sight when decanting the ethanol!

If I use isopropanol, I avoid cold temperatures because of the excess salt that usually comes down with it. If I want to increase the yields precipitated, I prefer to leave it incubating at room temperature longer vs. chilling the sample. When the DNA is pelleted, the pellet is sometimes more difficult to see compared to the ethanol pellet. It can be clear and glassy. Make sure, again, to note the side of the tube where the pellet should be. Look for it before decanting the isopropanol and 70% ethanol wash. After washing with ethanol, the pellet becomes visible and white. I always make sure it doesn’t slip off the side of the tube wall before decanting the supernatant. Allow the tube to drain upside down for a few minutes and then air dry or speed vac dry (5 minutes is enough) and then resuspend in buffer.

Finally, for dry DNA pellets, heating the sample in buffer at 50-60C will help the DNA dissolve faster and won’t damage the DNA. For RNA, heating can be used too (in water) at temps around 42C.  Overdried DNA and RNA will take longer to dissolve so make sure not to speed vac for too long.

So now you know the difference between ethanol and isopropanol and when to use which. If you have any questions, or anything to add, please drop a comment in below.

About the author

Suzanne Kennedy

Suzanne is Director of R&D at Mo Bio Laboratories in California, and the author of their blog, The Culture Dish. She has a PhD in Microbiology and Immunology from Virginia Commonwealth University.

To enable tagging you will need to register on Bitesize Bio. We're sorry for the inconvenience, but it's free, only takes a few seconds, and it will enable you to view our seminars for free, ask questions from the professional community, and take part in the lively community of Bitesize Bio

It’s that time of year again. The time when you have to fit in buying holiday gifts among the hundred other things you need to do at work and at home. Coming up with cool ideas or the “perfect gift” is a challenge for busy people with too much on their plate already.

To help our readers with their holiday shopping, here is a list of unique gift ideas for science geeks like us.  Whether you are looking for a  gift for a co-worker, a friends or you are looking for things you might like so you can drop hints to your signigicant others these ideas will help you hit the spot. If  you come up with some other great gift ideas, please share them by dropping us a comment!

So here are our gift ideas for science geeks:

Things to read

1. Cold Spring Harbor Labs Press’ library of novels, technical guides and text books can’t be beaten. A gift certificate to purchase from CSHLP would be a treat for any scientist.

2. A magazine subscription is a great gift that keeps on giving throughout the year! Amazon has a great selection of science-related magazine subscriptions for everything from Scientific American and the Smithsonian, right through to Wired.

3. Kindle Wireless Reading Device makes reading all of your important documents and novels easy; It holds up to 1500 books and can be read for up to 1 week on a single charge. So it’s perfect if you want to buy for someone who always has their arms full of papers or books.  Of course, this item is very expensive so it might be better as a group gift or maybe a nice present for the significant other in your life.

Things to wear

4. You can never go wrong with a t-shirt as a gift. Check out some of these fun and interesting t-shirts at everydayheroshirts.com.

5. If you are looking for a gift for a scientist who likes to stand out from the crowd, these specially commissioned cow- and snow leopard-print lab coats from The Lab Rat could be just the thing.

Things to make the lab look nice

6. While searching on Amazon.com I came across this cool selection of inspirational Einstein posters. Posters are a great way to spiff up the office or lab and so why not with the scientific genius!

7. One of my favorite gifts from a co-worker was a plant. Nothing brightens your office or lab space or desk like a potted plant and they last forever (or at least as long as you take care of them).

Cool geeky things

8. For the collectors on your list, fossils are a one of a kind gift. Check out a large selection of fossils at Fossil Mall. These range in price but are mostly expensive but are sure to be appreciated by anyone interested in archeology and prehistoric life.

9. Another unusual gift along the same lines is the Meteorite jewellery pendant from Science Mall USA.

10. And for your international friends or people who travel, take a look at some of these cool passport holders and laptop cases from Levenger.com.

Experiences

11. If you’ve got $5000 to spend, the ultimate gift could be a zero gravity flight. It might be expensive, but you can sooth your conscience with the fact that a ZERO-G keychain is included in the price.

12. Always wanted to be a writer but never feel like you have been given a chance? Bitesize Bio offers a free science writer experience to anyone with experience in the lab and the ability to string a few sentences together. Get the satisfaction of crafting your article, the fun of interacting with the Bitesize Bio staff as your article is polished and beautified and the thrill of knowing that your creation will be read by tens of thousands of scientists for years to come. This could be a gift to yourself or to someone you know. Drop us a line to get started! You won’t regret it. (ok, I’m kidding – you can do this at any time of course!)

Food and drink

13. Food is always a good gift for any hard-up graduate student so how about a gift card to Trader Joes (if you have one in the area) or gift certificates to your local favorite pub?

14. Or if your giftee is a caffeine addict, how about giving them a home-made gift voucher, offering to pay for the coffees each coffee break for a month?

Time – not every gift has to cost money

15. For friends who have a hectic family life, you can offer to take the weight off them for a while by doing dishes for the next month or free baby-sitting so they can have an evening out with their spouse. Who wouldn’t love that gift?

16. Or for people without families you could offer to make their life easier in the lab by looking after their cells or doing all of their minipreps for a month.

Gifts for others

17. Another option for the person who has everything is to buy a charitable gift in their name. Whether you (or they) would prefer to donate to Oxfam, the WWF or something completely different, a quick Google search will help you find a place to buy in their name.

18. And on a similar note, a worthy mention goes to the Toys for Tots campaign. Make a donation to them and they will use it to buy a gift for a kid who wouldn’t otherwise receive one this year. You could do this in lieu of a gift for someone else (or you could just do it anyway! ).

So those are my ideas, but whatever you buy, remember, it really is the thought that counts! Don’t forget to leave your gift ideas for scientists under this tree article before you leave.