Administrative note: RSS Feed
In my usual fashion, I plunged into this after doing all of 1 minute of homework. So I really didn't know what I was doing. Several people have suggested I needed RSS feed, but I couldn't find that at the pet store :-) Seriously, here it is (I assumed RSS came with things -- I have some learnin' to do!)
http://feeds.feedburner.com/OmicsOmics
If anyone is willing to comment, how gauche is it to host Google AdSense ads on a blog? With a little luck, I could use the proceeds to park a Solexa 1G in my garage, right? :-)
Tuesday, October 31, 2006
Monday, October 30, 2006
You can't always get what (samples) you want.
A key problem in omics research in medical research is getting the samples you need.
When I was an undergraduate, I had a fuzzy notion of a scheme for personalized medicine. Some analyzer would take a sample of what ailed you, look at it, and then generate a vial of customized antisense medicine that your doc would inject into you. I drove the pre-med in the lab nuts with my enthusiasm for it.
In graduate school, the analyzer became more clear: expression profiling. Look at the mRNA profile, figure out the disease, and voila, you are cured.
Fast forward to the latter part of my Millennium tenure. Rude surprise: you can rarely get the samples you want.
Most of my later work at Millennium was around cancer, originally because that is the research area I gravitated to & later because that was the one research area left (corporate evolution can be brutal!). Getting cancer samples turns out to be decidedly non-trivial.
If you are working in leukemia or related diseases (such as myeloproliferative syndromes), then things aren't bad. Your target tissue is floating around in the bloodstream & can be gotten with an ordinary blood draw. Patients in our society have been conditioned to expect lots of needle sticks, so this isn't hard.
For multiple myeloma and some lymphomas, you can go into the bone marrow. I'm a needlephobe, so the idea of a needle that crunches on the way in is decidedly unpleasant & sounds painful, and apparently is. Patients will do this infrequently, but not daily.
For a lot of solid tumors and other lymphomas, good luck -- particularly with recurrent disease. The tumors are hidden away (which is why they are often deadly) and quite small (if detected early). In many cases, getting a biopsy is surgical, painful, and perhaps significantly dangerous. You might get one sample; repeat visits are generally out of the question. Melanomas are one possible exception, but only for the primary lesion and not the metastases hiding everywhere.
This has significant implications. For a lot of studies, you would like to watch things over time. For example, what does the expression profile look like before and after drug treatment? How long does it take a pharmacodynamic protein marker to come up and what does its decay look like? Without multiple samples, these studies just can't happen.
Worse, what comes out may not be any good. Surgeons are in the business of saving lives, not going prospecting. Traditional practice is to cut first, then put away the samples after the patient is in recovery. But RNA & protein translational states are fragile, so if you don't pop the sample in liquid nitrogen immediately your sample may go downhill in a hurry. Multiple papers have reported finding expression signatures relating more to time-on-benchtop then any pathological state. It often takes dedicated personnel to perform this -- personnel the surgeons would rather not have 'in their way' (I've heard this first-hand from someone who used to be the sample grabber). A dirty not-so-secret in the business is that fresh frozen tissue just isn't practical for routine practice; you have to go with something else.
That is going to mean you go with several less palatable, but more available, options. One is to develop techniques to look at paraffin-embedded sections, which are the standard way of storing pathology samples. There are gazillions of such blocks sitting in hospitals, tempting the researchers. But, most of those sat on benchtops for uncontrolled time periods, so there may be some significant noise. Another is to try to fish the tiny number of tumor cells (or DNA) out of the bloodstream or perhaps an accessible fluid from the correct site (mucous from the lung; nipple aspirate for breast cancer). Or, you try to find markers in the blood or skin -- not where you are trying to treat, but easy to get to.
Whether these will work depends on what you are really looking for. For a predictive marker, it seems plausible that shed DNA or an old block might work. On the other hand, for a pharmacodynamic marker these are useless. A good PD marker allows you to measure whether your drug is hitting the target in vivo and at the correct site, and only by getting the real deal is that going to truly work. By necessity some studies use accessible non-tumor tissue, such as a skin punch or peripheral white blood cells, to at least see if the target is being hit somewhere. But that doesn't answer the question of whether the drug is getting to the tumor, a critical question. And many studies still use the traditional oncology PD marker of whether you are starting to destroy the patient's blood forming system.
At ASCO this summer, one speaker in a glioma section exhorted that a central repository for glioma samples must be imposed on the community, with a central authority determining who could do what experiments on which samples. That sort of extreme rationing shows how precious these samples are.
The scarcity of such samples also underlines why sensitive approaches, such as the nanowestern, are so critical. With small sample requirements, you might be able to go with fine needle biopsys rather than surgical biopsies, or be able to take lots of looks at the same sample (for different analytes).
Of course, things could be worse. What if you go to the trouble of getting a good sample, but then you look in the wrong place in that sample? Well, that's a post for another day.
A key problem in omics research in medical research is getting the samples you need.
When I was an undergraduate, I had a fuzzy notion of a scheme for personalized medicine. Some analyzer would take a sample of what ailed you, look at it, and then generate a vial of customized antisense medicine that your doc would inject into you. I drove the pre-med in the lab nuts with my enthusiasm for it.
In graduate school, the analyzer became more clear: expression profiling. Look at the mRNA profile, figure out the disease, and voila, you are cured.
Fast forward to the latter part of my Millennium tenure. Rude surprise: you can rarely get the samples you want.
Most of my later work at Millennium was around cancer, originally because that is the research area I gravitated to & later because that was the one research area left (corporate evolution can be brutal!). Getting cancer samples turns out to be decidedly non-trivial.
If you are working in leukemia or related diseases (such as myeloproliferative syndromes), then things aren't bad. Your target tissue is floating around in the bloodstream & can be gotten with an ordinary blood draw. Patients in our society have been conditioned to expect lots of needle sticks, so this isn't hard.
For multiple myeloma and some lymphomas, you can go into the bone marrow. I'm a needlephobe, so the idea of a needle that crunches on the way in is decidedly unpleasant & sounds painful, and apparently is. Patients will do this infrequently, but not daily.
For a lot of solid tumors and other lymphomas, good luck -- particularly with recurrent disease. The tumors are hidden away (which is why they are often deadly) and quite small (if detected early). In many cases, getting a biopsy is surgical, painful, and perhaps significantly dangerous. You might get one sample; repeat visits are generally out of the question. Melanomas are one possible exception, but only for the primary lesion and not the metastases hiding everywhere.
This has significant implications. For a lot of studies, you would like to watch things over time. For example, what does the expression profile look like before and after drug treatment? How long does it take a pharmacodynamic protein marker to come up and what does its decay look like? Without multiple samples, these studies just can't happen.
Worse, what comes out may not be any good. Surgeons are in the business of saving lives, not going prospecting. Traditional practice is to cut first, then put away the samples after the patient is in recovery. But RNA & protein translational states are fragile, so if you don't pop the sample in liquid nitrogen immediately your sample may go downhill in a hurry. Multiple papers have reported finding expression signatures relating more to time-on-benchtop then any pathological state. It often takes dedicated personnel to perform this -- personnel the surgeons would rather not have 'in their way' (I've heard this first-hand from someone who used to be the sample grabber). A dirty not-so-secret in the business is that fresh frozen tissue just isn't practical for routine practice; you have to go with something else.
That is going to mean you go with several less palatable, but more available, options. One is to develop techniques to look at paraffin-embedded sections, which are the standard way of storing pathology samples. There are gazillions of such blocks sitting in hospitals, tempting the researchers. But, most of those sat on benchtops for uncontrolled time periods, so there may be some significant noise. Another is to try to fish the tiny number of tumor cells (or DNA) out of the bloodstream or perhaps an accessible fluid from the correct site (mucous from the lung; nipple aspirate for breast cancer). Or, you try to find markers in the blood or skin -- not where you are trying to treat, but easy to get to.
Whether these will work depends on what you are really looking for. For a predictive marker, it seems plausible that shed DNA or an old block might work. On the other hand, for a pharmacodynamic marker these are useless. A good PD marker allows you to measure whether your drug is hitting the target in vivo and at the correct site, and only by getting the real deal is that going to truly work. By necessity some studies use accessible non-tumor tissue, such as a skin punch or peripheral white blood cells, to at least see if the target is being hit somewhere. But that doesn't answer the question of whether the drug is getting to the tumor, a critical question. And many studies still use the traditional oncology PD marker of whether you are starting to destroy the patient's blood forming system.
At ASCO this summer, one speaker in a glioma section exhorted that a central repository for glioma samples must be imposed on the community, with a central authority determining who could do what experiments on which samples. That sort of extreme rationing shows how precious these samples are.
The scarcity of such samples also underlines why sensitive approaches, such as the nanowestern, are so critical. With small sample requirements, you might be able to go with fine needle biopsys rather than surgical biopsies, or be able to take lots of looks at the same sample (for different analytes).
Of course, things could be worse. What if you go to the trouble of getting a good sample, but then you look in the wrong place in that sample? Well, that's a post for another day.
Sunday, October 29, 2006
Nanowesterns: The future of signal transduction research?
Western blots are a workhorse of biology. When everything goes right, they allow for interrogating the state & quantity of a protein in a cellular system. They can be exquisitely sensitive and specific; Western blot assays have long been used as the definitive test for a number of medical conditions, particularly HIV infection. Given the right antibody, you can detect anything, including miniscule amounts of phosphorylated proteins. And, to a first approximation, they are quantitative.
A Western blot involves several steps. First, the samples of interest are placed in a denaturing buffer, causing the proteins to unfold & disaggregate. The unfolding is performed by large quantities of detergent (primarily SDS, which also shows up in your toothpaste, laundry detergent, dishwashing liquid, etc -- the stuff is ubiquitous) and the disaggregation is assisted by some sort of sulfhydryl compound to destroy disulfide bridges. Such compounds are uniformly smelly, except to a lucky few (I had a graduate school colleague who was smell-blind for them).
Now the samples are loaded on an SDS-PAGE gel, which uses electricity to separate the proteins by size -- approximately. In theory. Once they are separated, the proteins are transferred to a membrane by osmosis or electrophoresis perpendicular to the first direction. The extra protein binding sites on the membrane are then blocked, often with Carnation non-fat dry milk (I kid you not; the stuff is cheap & works). An antibody for the target of interest is added, and then an antibody to detect the first antibody; this one carries a label of some sort. Occasionally it is a third antibody which detects the 2nd (which bound the first) -- each level can enhance sensitivity. The appropriate detection chemistry is run & voila! You have a Western blot. Between all the steps after the blocking are lots of washes to remove excess reagents.
The beauty of a Western is that the technology is pretty cheap & simple -- I did a bunch of Western's in my senior thesis in a lab that ran on a shoestring budget -- and I'm all thumbs in the lab. The truly amazing part is that Westerns today are run pretty much the same way. You might buy pre-poured gels, but the basics are all the same.
The problems are legion.
First, this is a decidedly low-throughput assay scheme -- typical gels have maybe two dozen lanes for running. This is one reason it is used as a confirmatory test for HIV and other infections; large scale testing is out of the question.
Second, it is very labor intensive. Setting up the transfer from gel to membrane is inherently a manual process, but somewhat surprisingly it still seems uncommon to automate the later steps or even the washing. During a short lived Western blot process improvement project I initiated, I discovered that the folks running the blots both disliked the washing but also found it a social activity -- everyone is doing something mindless, so there is time to talk (simple fly pushing in a Drosophila lab is similar; the lab I was in almost always had NPR going in the background).
Third, they can require a lot of tuning. Different extraction ("lysis") buffers for the initial extraction, different gel or running conditions, different membranes, antibody dilutions, etc. -- these are all variables one can play with on the blot. Some rules of thumb are out there based on the location of the protein or how greasy it is, but it is clearly more art & lore than science. Some proteins never seem to work. Many result in big messes -- which is another advantage of Westerns, as you have the electrophoretic separation perhaps parsing the mixture into uninteresting bands & the one you want -- which may well be much fainter than the junk. And, of course, the gels don't always run the right way. Too hot -- trouble. Not poured evenly -- trouble. And please don't drop them on the floor!
Some of the trouble comes from the antibodies, but that is easily a topic for another time. But most is inherent in the Western scheme (no, there was no Dr. Western -- but there was a Dr. Southern and the other compass blots are plays on that).
But they are still extremely useful. Some folks have tried to push the envelope within the boundaries of a conventional Western. Perhaps the best example of this has been commercialized at Kinexus, which has pushed multiplex Westerns to amazing limits. They work carefully to identify sets of antibodies which will not interfere with each other & which also generate non-overlapping bands. One way to think about this is a really good Western antibody generates a single band in the same spot on the gel -- which means the rest of the blot is wasted data. Kinexus tries to maximize the amount of data from one gel. But this is a lot of work.
A new publication (free!) describes an approach that has a bit in common with the Western, but in many ways is altogether a different beast. The work is done by a startup called Cell Biosciences.
The slab gel is replaced by capillaries -- easy to control on the thermal side. SDS-PAGE is replaced by isoelectric focusing (IEF). Instead of blotting to a membrane, the separated proteins are locked onto the wall of the capillary. But other than everything being different, it's a Western!
Isoelectric focusing is a technique for electrophoretic separation of proteins. Instead of size, which SDS-PAGE sorts on, IEF uses protein charge. Each protein contains some amino acids which have positive charge and some with negative charge, plus postively & negatively charged ends. Post-translational modifications can further stir the pot: phosphorylation adds two negative charges per phosphate, and something big like ubiquitin tacks on a complex mess. On the other end, some modifications, such as acetylation, may replace a charged group with an uncharged one. In a gel with a pH gradient & subject to an electric field, the proteins will migrate to the pH where they have no charge -- the positively ionized and negatively ionized groups are in perfect balance.
An advantage of this pointed out in the nanowestern paper is that you can load a lot more sample on a gel. For a size separation, only a narrow band of sample can enter the gel because the separation is based on different sized proteins traveling at different speeds. Because IEF is an equilibrium method, you can actually fill the entire capillary with sample and then apply the electric field. This has important sensitivity implications.
The paper also describes a whole apparatus for automating the whole shebang; quite a contrast from an ordinary western. The model described runs only a dozen capillaries, but modern DNA sequencers routinely run hundreds simultaneously so there is plenty of room to grow. Each capillary detects one analyte for one sample, so with hundreds you could process hundreds of samples or detect hundreds of analytes, or some interesting middle ground.
Capillaries are also intriguing because they are at the heart of many lab-on-a-chip schemes. This paper might suggest the notion of a multi-analyte integrated chip.
The paper also describes using multiple fluorescent peptides as internal standards. These are synthesized in the opposite handedness as natural peptides, this doesn't change their IEF properties, but does make them unpalatable to proteases that might be present in the sample (though in general you use cocktails of inhibitors to prevent those proteases from attacking your sample).
The authors describe using a single antibody to assess the phosphorylation states of two related proteins, ERK1 and ERK2. Such determinations can be challenging on a Western if the two run closely with each other. In an SDS-PAGE gel, the behavior of phosphorylated proteins is maddening -- sometimes they run with the unphosphorylated form and sometimes they form new bands (a "phosphoshift"). Murphy's law rules; whichever behavior you don't want is the one you get! With IEF, phosphorylation should always give a strong phosphoshift, as a weakly ionizable side chain (hydroxyl on a Ser, Thr or Tyr) is replaced with a strongly acidic phosphate.
The paper describes using their system with a few proteins. That's a good start, but I suspect most people will want to see more. A lot more. And with other modifications. Ubiquitination would be particularly interesting, both because it is a big modification and because ubiquitin chains are often form. Presumably this will lead to a laddering effect. Also interesting to look at are more complicated phosphorylation systems than the ones examined here, with tens of phosphorylation sites rather than a handful. A reasonable guess is that the approach will still count sites, but if you want to distinguish them, which generally you will, you will still need specific antibodies for each site.
One last plus of their scheme, which they place right in the title & is the source of the nano moniker. The system is very sensitive, at least with the one analyte tested. This is important for rare or small samples (more on samples in another post). They even claim they might be able to push it from 25 cells down to 1 cell. If this is true, or even if 25 cells is achieved consistently with many antibodies, this will be an impressive feat & make this technique very attractive for signal transduction research.
Western blots are a workhorse of biology. When everything goes right, they allow for interrogating the state & quantity of a protein in a cellular system. They can be exquisitely sensitive and specific; Western blot assays have long been used as the definitive test for a number of medical conditions, particularly HIV infection. Given the right antibody, you can detect anything, including miniscule amounts of phosphorylated proteins. And, to a first approximation, they are quantitative.
A Western blot involves several steps. First, the samples of interest are placed in a denaturing buffer, causing the proteins to unfold & disaggregate. The unfolding is performed by large quantities of detergent (primarily SDS, which also shows up in your toothpaste, laundry detergent, dishwashing liquid, etc -- the stuff is ubiquitous) and the disaggregation is assisted by some sort of sulfhydryl compound to destroy disulfide bridges. Such compounds are uniformly smelly, except to a lucky few (I had a graduate school colleague who was smell-blind for them).
Now the samples are loaded on an SDS-PAGE gel, which uses electricity to separate the proteins by size -- approximately. In theory. Once they are separated, the proteins are transferred to a membrane by osmosis or electrophoresis perpendicular to the first direction. The extra protein binding sites on the membrane are then blocked, often with Carnation non-fat dry milk (I kid you not; the stuff is cheap & works). An antibody for the target of interest is added, and then an antibody to detect the first antibody; this one carries a label of some sort. Occasionally it is a third antibody which detects the 2nd (which bound the first) -- each level can enhance sensitivity. The appropriate detection chemistry is run & voila! You have a Western blot. Between all the steps after the blocking are lots of washes to remove excess reagents.
The beauty of a Western is that the technology is pretty cheap & simple -- I did a bunch of Western's in my senior thesis in a lab that ran on a shoestring budget -- and I'm all thumbs in the lab. The truly amazing part is that Westerns today are run pretty much the same way. You might buy pre-poured gels, but the basics are all the same.
The problems are legion.
First, this is a decidedly low-throughput assay scheme -- typical gels have maybe two dozen lanes for running. This is one reason it is used as a confirmatory test for HIV and other infections; large scale testing is out of the question.
Second, it is very labor intensive. Setting up the transfer from gel to membrane is inherently a manual process, but somewhat surprisingly it still seems uncommon to automate the later steps or even the washing. During a short lived Western blot process improvement project I initiated, I discovered that the folks running the blots both disliked the washing but also found it a social activity -- everyone is doing something mindless, so there is time to talk (simple fly pushing in a Drosophila lab is similar; the lab I was in almost always had NPR going in the background).
Third, they can require a lot of tuning. Different extraction ("lysis") buffers for the initial extraction, different gel or running conditions, different membranes, antibody dilutions, etc. -- these are all variables one can play with on the blot. Some rules of thumb are out there based on the location of the protein or how greasy it is, but it is clearly more art & lore than science. Some proteins never seem to work. Many result in big messes -- which is another advantage of Westerns, as you have the electrophoretic separation perhaps parsing the mixture into uninteresting bands & the one you want -- which may well be much fainter than the junk. And, of course, the gels don't always run the right way. Too hot -- trouble. Not poured evenly -- trouble. And please don't drop them on the floor!
Some of the trouble comes from the antibodies, but that is easily a topic for another time. But most is inherent in the Western scheme (no, there was no Dr. Western -- but there was a Dr. Southern and the other compass blots are plays on that).
But they are still extremely useful. Some folks have tried to push the envelope within the boundaries of a conventional Western. Perhaps the best example of this has been commercialized at Kinexus, which has pushed multiplex Westerns to amazing limits. They work carefully to identify sets of antibodies which will not interfere with each other & which also generate non-overlapping bands. One way to think about this is a really good Western antibody generates a single band in the same spot on the gel -- which means the rest of the blot is wasted data. Kinexus tries to maximize the amount of data from one gel. But this is a lot of work.
A new publication (free!) describes an approach that has a bit in common with the Western, but in many ways is altogether a different beast. The work is done by a startup called Cell Biosciences.
The slab gel is replaced by capillaries -- easy to control on the thermal side. SDS-PAGE is replaced by isoelectric focusing (IEF). Instead of blotting to a membrane, the separated proteins are locked onto the wall of the capillary. But other than everything being different, it's a Western!
Isoelectric focusing is a technique for electrophoretic separation of proteins. Instead of size, which SDS-PAGE sorts on, IEF uses protein charge. Each protein contains some amino acids which have positive charge and some with negative charge, plus postively & negatively charged ends. Post-translational modifications can further stir the pot: phosphorylation adds two negative charges per phosphate, and something big like ubiquitin tacks on a complex mess. On the other end, some modifications, such as acetylation, may replace a charged group with an uncharged one. In a gel with a pH gradient & subject to an electric field, the proteins will migrate to the pH where they have no charge -- the positively ionized and negatively ionized groups are in perfect balance.
An advantage of this pointed out in the nanowestern paper is that you can load a lot more sample on a gel. For a size separation, only a narrow band of sample can enter the gel because the separation is based on different sized proteins traveling at different speeds. Because IEF is an equilibrium method, you can actually fill the entire capillary with sample and then apply the electric field. This has important sensitivity implications.
The paper also describes a whole apparatus for automating the whole shebang; quite a contrast from an ordinary western. The model described runs only a dozen capillaries, but modern DNA sequencers routinely run hundreds simultaneously so there is plenty of room to grow. Each capillary detects one analyte for one sample, so with hundreds you could process hundreds of samples or detect hundreds of analytes, or some interesting middle ground.
Capillaries are also intriguing because they are at the heart of many lab-on-a-chip schemes. This paper might suggest the notion of a multi-analyte integrated chip.
The paper also describes using multiple fluorescent peptides as internal standards. These are synthesized in the opposite handedness as natural peptides, this doesn't change their IEF properties, but does make them unpalatable to proteases that might be present in the sample (though in general you use cocktails of inhibitors to prevent those proteases from attacking your sample).
The authors describe using a single antibody to assess the phosphorylation states of two related proteins, ERK1 and ERK2. Such determinations can be challenging on a Western if the two run closely with each other. In an SDS-PAGE gel, the behavior of phosphorylated proteins is maddening -- sometimes they run with the unphosphorylated form and sometimes they form new bands (a "phosphoshift"). Murphy's law rules; whichever behavior you don't want is the one you get! With IEF, phosphorylation should always give a strong phosphoshift, as a weakly ionizable side chain (hydroxyl on a Ser, Thr or Tyr) is replaced with a strongly acidic phosphate.
The paper describes using their system with a few proteins. That's a good start, but I suspect most people will want to see more. A lot more. And with other modifications. Ubiquitination would be particularly interesting, both because it is a big modification and because ubiquitin chains are often form. Presumably this will lead to a laddering effect. Also interesting to look at are more complicated phosphorylation systems than the ones examined here, with tens of phosphorylation sites rather than a handful. A reasonable guess is that the approach will still count sites, but if you want to distinguish them, which generally you will, you will still need specific antibodies for each site.
One last plus of their scheme, which they place right in the title & is the source of the nano moniker. The system is very sensitive, at least with the one analyte tested. This is important for rare or small samples (more on samples in another post). They even claim they might be able to push it from 25 cells down to 1 cell. If this is true, or even if 25 cells is achieved consistently with many antibodies, this will be an impressive feat & make this technique very attractive for signal transduction research.
Saturday, October 28, 2006
Starting out
Okay, here goes nothing. Millennium Pharmaceuticals just sloughed off its genomics assets, which included me. So I have some time on my hands to try something new.
I've spent 10 years with Millennium, and prior to that I was trained in the labs of two pioneers in genomics, Wally Gilbert & George Church. I'm a biologist by training, a computational biologist by preference, and an omics-fan by heart. At Millennium I played with or thought about about every omics technology there is, from DNA sequencing to expression profiling to proteomics. I won't claim I am an expert on all of them; just that I have ideas (perhaps deluded) about how they might be used.
I'll try to make a few initial promises & a few requests of you, the readers I haven't found yet. First, I will try to make my biases clear. For example, I have a hard time being objective about stuff coming from the Church lab -- my heart will long be there.
Second, I won't be a touter or a trasher. I'll try to be fair. Other than my former employer, I don't have a financial stake in anything; all other investments are in broad index funds. If my next job or some consulting gig is at a technology company, then I'll try to be open about that or avoid subjects that present a conflict-of-interest. Please don't use this blog to invest or ask me for stock tips -- I confidently predicted the Tigers would sweep this year's World Series.
Third, I'll try to stay somewhat within my expertise. If you catch me saying something dumb, please point it out. But most of the interesting stuff requires a stretch, and I strongly believe that the best results come from vigorously hashing lots of half-baked ideas -- eventually you find the ones worth sticking back in the oven.
Fourth, I'll try to keep this focused. Minimal straying off into politics or beating up on pseudoscience. It's not that I don't care about these things, it's just I need the practice keeping something focused.
Fifth, this won't be a blow-by-blow of my job search. I'd love to get leads on compuational biology positions in Boston/Cambridge or the burbs (particularly north/west of the city), but I'm not going to go blow-by-blow on my interviews. That's not what this is for.
Sixth, this won't be a place to routinely trash or praise my former employer. Millennium was an exciting place where I learned a lot, but we made plenty of mistakes too. I'll be referring to my time there frequently, but I'll try to keep some balance.
The name? Well, I played with several. I have a fondness for silliness, and the Little Caesars ads fit that. It fit the subject matter & didn't seem to be taken already. Now all I need is a picture of a little DNA helix with a laurel wreath & a spear...
I've spent 10 years with Millennium, and prior to that I was trained in the labs of two pioneers in genomics, Wally Gilbert & George Church. I'm a biologist by training, a computational biologist by preference, and an omics-fan by heart. At Millennium I played with or thought about about every omics technology there is, from DNA sequencing to expression profiling to proteomics. I won't claim I am an expert on all of them; just that I have ideas (perhaps deluded) about how they might be used.
I'll try to make a few initial promises & a few requests of you, the readers I haven't found yet. First, I will try to make my biases clear. For example, I have a hard time being objective about stuff coming from the Church lab -- my heart will long be there.
Second, I won't be a touter or a trasher. I'll try to be fair. Other than my former employer, I don't have a financial stake in anything; all other investments are in broad index funds. If my next job or some consulting gig is at a technology company, then I'll try to be open about that or avoid subjects that present a conflict-of-interest. Please don't use this blog to invest or ask me for stock tips -- I confidently predicted the Tigers would sweep this year's World Series.
Third, I'll try to stay somewhat within my expertise. If you catch me saying something dumb, please point it out. But most of the interesting stuff requires a stretch, and I strongly believe that the best results come from vigorously hashing lots of half-baked ideas -- eventually you find the ones worth sticking back in the oven.
Fourth, I'll try to keep this focused. Minimal straying off into politics or beating up on pseudoscience. It's not that I don't care about these things, it's just I need the practice keeping something focused.
Fifth, this won't be a blow-by-blow of my job search. I'd love to get leads on compuational biology positions in Boston/Cambridge or the burbs (particularly north/west of the city), but I'm not going to go blow-by-blow on my interviews. That's not what this is for.
Sixth, this won't be a place to routinely trash or praise my former employer. Millennium was an exciting place where I learned a lot, but we made plenty of mistakes too. I'll be referring to my time there frequently, but I'll try to keep some balance.
The name? Well, I played with several. I have a fondness for silliness, and the Little Caesars ads fit that. It fit the subject matter & didn't seem to be taken already. Now all I need is a picture of a little DNA helix with a laurel wreath & a spear...
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