Monday, February 7, 2011
A 19.2V Drill with every sorter purchase?
As you've probably heard, Beckman Coulter put itself up for sale last year, and now it looks like they will be purchased by Washington based Danaher Corporation. Danaher is a conglomerate that owns the Craftsman hand tool brand as well as businesses in electronic testing equipment, dental equipment, and monitoring products. They also own the microscopy company Leica. What does this mean for the flow cytometry world? Probably not much, but the extra capital available from Danaher could only help R&D for flow cytometry, right? Time will tell. It seems like the MoFlo just can't find a stable home. Ever since Cytomation was acquired by Dako, the MoFlo has been passed around like a plate of crudités at a 6 year old's birthday party. My advice to Danaher and those making the transition from Beckman Coulter - forget everything you think you know about flow cytometry, come talk to the folks in trenches, and design a new instrument that's more than a "me-too" product. Good luck! Here's Beckman-Coulter's statement. And here's what Danaher has to say.
Friday, January 7, 2011
The Beckman Coulter Gallios and Translational Research at UCFlow
Have you ever had a frantic MD fellow rush into the lab with a rack full of tubes saying, "A patient just showed up and I have these samples that I need to run now. Is there any instrument time available?" A couple of years ago, I would have said no, but this scenario has become more and more common around here. As the lines between the clinic and the research bench become ever more blurred, the needs of the community in a research medical center begin to expand. The major variable in this whole arena is a fickle creature we like to call a human being. You see, unlike mice, they don't get sick when you tell them to and you can't force them to fit into your schedule, so you have no choice but to modify your service to accommodate the unpredictable nature of clinical research. The perceived lack of access to the core by clinical researchers has also been the driving force behind individual researchers' desire to work outside of the core and invest in their own instrumentation. In principle, I don't really have a problem with this, but as a business model in a University, I think it's very inefficient. We have spent years perfecting our craft in the flow lab. In fact, we possess a collective 25+ years of experience operating, maintaining, and troubleshooting flow cytometers and sorters. It's difficult to see why someone would want to side-step all that knowledge. But, I digress...
At the same time we were scratching our heads as to how we could offer this group of clinical researchers greater access to flow instrumentation while not taking away capacity from our large and active group of basic researchers, we were evaluating and testing an instrument called the Gallios from Beckman Coulter. The Gallios is a fine piece of hardware. It has some of the things you'd expect of an analyzer from Coulter, flashing lights (like the FC500), a carousel loader, a fairly locked-down box. But it also has some new/unexpected things. They took a cue from the success BD has had marketing the "Octagon" and came up with their own design called the "Boulevard." It basically serves the same purpose; bounce light off filters, don't transmit light through a bunch of filters. They deliver light to the Boulevard via a fiber cable coupled to a pinhole for the appropriate laser - pretty much the same as a BD instruments. Another interesting optical component is the laser launch module. The solid state lasers shoot their beam into a steering tower that has motor controlled micrometers which allows for remote alignment. Laser light to the flow cell is delivered in air, not fiber, as to maximize energy at the point of illumination. These two things make the Gallios pretty much optically on par with an LSRII. There's a unique FSC detector that tries to look at different angles of refraction to better resolve small particles, but since I don't care too much about that, I'll skip it. The thing that sets the Gallios apart from the DiVa setup is in fact the electronics. I won't attempt to explain the architecture of both platforms here, but will simply cut to the chase. More bits of resolution across 4 or 5 log decades leads to better resolution of dimly stained cells when comparing two instruments that are optically pretty similar. So, in my testing, I was able to resolve dim stuff from background better on the Gallios than on my LSRII, and the reason that's the case, in my opinion, is the higher resolution electronics on the Gallios (especially when comparing the 1st and 2nd decade of the scale). The last thing about the Gallios was its optical stability. Again, comparing it to our LSRII, which we tweak the alignment more frequently than we'd like to, the Gallios is rock solid. It probably compares pretty well to the optical stability of a FACSCanto-II (as I've heard from others who have one - I only have FACSCanto-As, which I don't like at all). But, even as of yesterday, about 10 months after install, the beads look exactly the same. I've never seen any of my instruments not need a little tweak of the alignment after 10 months of use. That was impressive.
So, I had an instrument that seemed to work really well for us, and I had a problem with capacity for last-minute clinical research use. Are you thinking what I'm thinking??? You got it, kill two birds with one stone (figuratively of course; I don't condone the practice of killing anything with stones). So, I wrote a proposal to the department to pitch the idea of buying the Gallios and opening it up to our clinical research group only. This would free them from worrying about having time booked in advance on one of our instruments. In fact, booking time on the instrument wouldn't even be allowed more than 48 hours in advance. Thankfully the department liked the idea, awarded us the money, and we're developing the usage plan now to offer this service to our Translational Research groups. To sweeten the deal a bit, we offered to expand our Drop-off service so that clincian/researchers who did not have a lab full of techs could simply drop the samples off to us and we would run them on the Gallios and give them back a preliminary analysis of the data. Win-Win-Win for all.
At the same time we were scratching our heads as to how we could offer this group of clinical researchers greater access to flow instrumentation while not taking away capacity from our large and active group of basic researchers, we were evaluating and testing an instrument called the Gallios from Beckman Coulter. The Gallios is a fine piece of hardware. It has some of the things you'd expect of an analyzer from Coulter, flashing lights (like the FC500), a carousel loader, a fairly locked-down box. But it also has some new/unexpected things. They took a cue from the success BD has had marketing the "Octagon" and came up with their own design called the "Boulevard." It basically serves the same purpose; bounce light off filters, don't transmit light through a bunch of filters. They deliver light to the Boulevard via a fiber cable coupled to a pinhole for the appropriate laser - pretty much the same as a BD instruments. Another interesting optical component is the laser launch module. The solid state lasers shoot their beam into a steering tower that has motor controlled micrometers which allows for remote alignment. Laser light to the flow cell is delivered in air, not fiber, as to maximize energy at the point of illumination. These two things make the Gallios pretty much optically on par with an LSRII. There's a unique FSC detector that tries to look at different angles of refraction to better resolve small particles, but since I don't care too much about that, I'll skip it. The thing that sets the Gallios apart from the DiVa setup is in fact the electronics. I won't attempt to explain the architecture of both platforms here, but will simply cut to the chase. More bits of resolution across 4 or 5 log decades leads to better resolution of dimly stained cells when comparing two instruments that are optically pretty similar. So, in my testing, I was able to resolve dim stuff from background better on the Gallios than on my LSRII, and the reason that's the case, in my opinion, is the higher resolution electronics on the Gallios (especially when comparing the 1st and 2nd decade of the scale). The last thing about the Gallios was its optical stability. Again, comparing it to our LSRII, which we tweak the alignment more frequently than we'd like to, the Gallios is rock solid. It probably compares pretty well to the optical stability of a FACSCanto-II (as I've heard from others who have one - I only have FACSCanto-As, which I don't like at all). But, even as of yesterday, about 10 months after install, the beads look exactly the same. I've never seen any of my instruments not need a little tweak of the alignment after 10 months of use. That was impressive.
So, I had an instrument that seemed to work really well for us, and I had a problem with capacity for last-minute clinical research use. Are you thinking what I'm thinking??? You got it, kill two birds with one stone (figuratively of course; I don't condone the practice of killing anything with stones). So, I wrote a proposal to the department to pitch the idea of buying the Gallios and opening it up to our clinical research group only. This would free them from worrying about having time booked in advance on one of our instruments. In fact, booking time on the instrument wouldn't even be allowed more than 48 hours in advance. Thankfully the department liked the idea, awarded us the money, and we're developing the usage plan now to offer this service to our Translational Research groups. To sweeten the deal a bit, we offered to expand our Drop-off service so that clincian/researchers who did not have a lab full of techs could simply drop the samples off to us and we would run them on the Gallios and give them back a preliminary analysis of the data. Win-Win-Win for all.
Tuesday, November 9, 2010
Compensation is infiltrating my dreams.
I've been composing this post in my head for a couple of weeks now, but have been too busy to sit down and write it, so it shouldn't really surprise me that it popped up in a dream. However, a new twist was added via my unconscious mind (which I'll get to later). So, the original post was all about how I've pretty much given up on compensating using cells, and if you're not using beads, then you're pretty much setting yourself up for compensation failure (unless of course you're using things like PI, or mCherry, or the like). I mean, the whole point of 'autocomp' is to take the subjectivity out of compensation, and using objective mathematics to correct for fluorescence spillover. However, every single time I've done autocomp using cells, it just doesn't look 'right' and I end up tweaking the values just a little bit. I've come to terms with this fact, and have pretty much settled with this sub-par situation. But, if you're trying to teach someone about compensation, and you introduce this 'autocomp' feature, it makes for a pretty awkward conversation when you then go on to say, "Well, just adjust the values a little bit until it looks right." So, I typically recommend people do their compensation with beads. For many of my users, the thing that prevents them from doing this is cost, or maybe a bit of skepticism in changing the ways they were taught to do their staining. The reasons why compensating using cells doesn't always work are many, but let me just outline a few for you here.
1. Insufficient frequencies of both positive and negative fraction to make a statistically significant regression of means. If in your stained cell sample, you only have a 0.1% positive fraction, the mean of that population in the spillover channel will not reach a high enough statistical significance until you collect millions of cells. No one is going to collect millions of cells on their single stain control. This also holds true when all your cells are positive for your single stain control, and you have a really low negative (or low) population.
2. Poor resolution of the positive fraction. Sometimes you will not have a clear positive population, so making a gate around the positive fraction for performing compensation is difficult. If you end up encircling some of the high autofluorescent cells that you mistakenly call positive, your compensation will surely be off.
3. Non-linearities at the extremes can lead to inaccurate compensation. If you're compensating using an unstained (or negative) fraction that is at the very low-end of the scale, or if your positive fraction is at the very high-end of the scale, you're likely using a data point in the non-linear range of the log scale. Since compensation algorithms are basically relying on the fact that your range of analysis is linear, you're going to run into lots of problems if you're using "unstained" cells as your low-data point, or really bright cells as your high data point. Side bar: Yes, I know, your comp control should be at least as bright as your sample staining, blah, blah, blah. However, the only reason why this is the case is because of non-linearities at the very high end of the scale. If all your staining fell within the linear portion of the scale (let's say 1.0 logs to 3.5 logs), then this isn't necessarily a problem. You can take any two points within that range, and create a regression line that will model the entire scale. No-scale is linear enough, especially at the extremes, so the 'rule' of a maximally bright comp control needs to be adhered to.
4. Mismatched autofluorescence between positive and negative. If I stained my leukocyte prep with a monocyte marker (CD14, for example). All my monocytes will be positive. For this single stained comp control, what should I use as my negative? Many people would simply use the negative lymphocytes or granulocytes, and many people would end up with a poor compensation matrix. For channels where autofluorescence is a factor (mostly the green/yellow detectors off the blue and lower laser lines), the positive fraction's autofluorescence should match the negative fraction's autofluorescence. This is, evidently only necessary when you're using cells for compensation, and you have a mixed cell-type sample.
So, there are certainly lots of pitfalls when using cells for compensation, which is why using beads is a good idea. To solve many of these issues, simply using an antibody capture bead at two fluorescence levels should do the trick. You'll notice I said two fluorescence levels, and not one positive and the 'blank' bead. Using the blank bead can lead us into issue #3 above, so I prefer to use the bead at a saturating level of antibody and maybe 100-fold less, to create a high and low peak. In the end the peaks will fall around the 3.5 decade range and 1.5 decade range. Use these peaks as your 'positive' and 'negative' values in your favorite autocomp program, and voila, perfect compensation. Of course, these beads are run at the appropriate voltage that is set up according to your cell type.
But, what about the twist? The twist is, that you don't need to only use beads as your capture matrix. You could use cells! I know, I know, I just went on and on about NOT using cells, now I'm telling you to use cells, but wait, let me explain. Take a thymus, get all your non-tandem antibodies in CD4, stain them at two concentrations, fix them, and stick them in the fridge. You now have ready-made compensation controls that are much cheaper than buying capture beads. Why thymus? They're the closest thing to beads; pretty much homogeneous, so we don't have to worry about autofluorescence mismatch, they're almost all CD4 positive, so that makes it easier to create two nice peaks, and you can get a boatload of them from a young mouse. On top of all this, we gain the ability to use other things besides antibodies. You could stain them with many of your dyes for a comp control, PI, DAPI, CFSE, etc... Something you can't do with beads. For tandems, I'd stick with capture beads.
Ok, there you have it. If you've made it this far reading through all my gibberish, let me know what you think.
1. Insufficient frequencies of both positive and negative fraction to make a statistically significant regression of means. If in your stained cell sample, you only have a 0.1% positive fraction, the mean of that population in the spillover channel will not reach a high enough statistical significance until you collect millions of cells. No one is going to collect millions of cells on their single stain control. This also holds true when all your cells are positive for your single stain control, and you have a really low negative (or low) population.
2. Poor resolution of the positive fraction. Sometimes you will not have a clear positive population, so making a gate around the positive fraction for performing compensation is difficult. If you end up encircling some of the high autofluorescent cells that you mistakenly call positive, your compensation will surely be off.
3. Non-linearities at the extremes can lead to inaccurate compensation. If you're compensating using an unstained (or negative) fraction that is at the very low-end of the scale, or if your positive fraction is at the very high-end of the scale, you're likely using a data point in the non-linear range of the log scale. Since compensation algorithms are basically relying on the fact that your range of analysis is linear, you're going to run into lots of problems if you're using "unstained" cells as your low-data point, or really bright cells as your high data point. Side bar: Yes, I know, your comp control should be at least as bright as your sample staining, blah, blah, blah. However, the only reason why this is the case is because of non-linearities at the very high end of the scale. If all your staining fell within the linear portion of the scale (let's say 1.0 logs to 3.5 logs), then this isn't necessarily a problem. You can take any two points within that range, and create a regression line that will model the entire scale. No-scale is linear enough, especially at the extremes, so the 'rule' of a maximally bright comp control needs to be adhered to.
4. Mismatched autofluorescence between positive and negative. If I stained my leukocyte prep with a monocyte marker (CD14, for example). All my monocytes will be positive. For this single stained comp control, what should I use as my negative? Many people would simply use the negative lymphocytes or granulocytes, and many people would end up with a poor compensation matrix. For channels where autofluorescence is a factor (mostly the green/yellow detectors off the blue and lower laser lines), the positive fraction's autofluorescence should match the negative fraction's autofluorescence. This is, evidently only necessary when you're using cells for compensation, and you have a mixed cell-type sample.
So, there are certainly lots of pitfalls when using cells for compensation, which is why using beads is a good idea. To solve many of these issues, simply using an antibody capture bead at two fluorescence levels should do the trick. You'll notice I said two fluorescence levels, and not one positive and the 'blank' bead. Using the blank bead can lead us into issue #3 above, so I prefer to use the bead at a saturating level of antibody and maybe 100-fold less, to create a high and low peak. In the end the peaks will fall around the 3.5 decade range and 1.5 decade range. Use these peaks as your 'positive' and 'negative' values in your favorite autocomp program, and voila, perfect compensation. Of course, these beads are run at the appropriate voltage that is set up according to your cell type.
But, what about the twist? The twist is, that you don't need to only use beads as your capture matrix. You could use cells! I know, I know, I just went on and on about NOT using cells, now I'm telling you to use cells, but wait, let me explain. Take a thymus, get all your non-tandem antibodies in CD4, stain them at two concentrations, fix them, and stick them in the fridge. You now have ready-made compensation controls that are much cheaper than buying capture beads. Why thymus? They're the closest thing to beads; pretty much homogeneous, so we don't have to worry about autofluorescence mismatch, they're almost all CD4 positive, so that makes it easier to create two nice peaks, and you can get a boatload of them from a young mouse. On top of all this, we gain the ability to use other things besides antibodies. You could stain them with many of your dyes for a comp control, PI, DAPI, CFSE, etc... Something you can't do with beads. For tandems, I'd stick with capture beads.
Ok, there you have it. If you've made it this far reading through all my gibberish, let me know what you think.
Friday, October 1, 2010
MoFlo Upgraded to XDP, plus a couple new laser lines.
Ah, the MoFlo - what a fine piece of craftsmanship! I started my relationship with the MoFlo (Formerly of Cytomation, Formerly of DakoCytomation, Formerly of Dako, Currently of Beckman-Coulter) in the year 2000. We had many great years together, but our relationship was getting a bit stale. You see, there was this fancy new gal in town call the Aria who lured me into her web of seduction with promises of 'turn-key' operation, and I bit! I soon realized however, that the grass isn't necessarily greener on the other side, and re-visited the rock-solid usability of the MoFlo. In recent years, the MoFlo started showing its age. I have to admit, part of the issue was a certain level of neglect and abuse on our part, but hey 10 years in instrument years is like 80 in people years. And so we came to a fork in the road, and as with most things in the technology area utilizing 20 year old components, we had to decide, pull the plug or pursue the upgrade path.
When I was contacted by the folks at Propel labs (who, evidently are a group of people from the original Cytomation company) that there was an upgrade path to the XDP electronics for the legacy MoFlo, I was thrilled. After about a year of begging for money from anyone that would listen to me, I finally secured the funding and was ready for the upgrade. So, why upgrade to XDP instead of buying a new sorter? Well, first of all, it was a financial thing. The cost of an upgrade is about 1/4th the cost of a new sorter. Secondly, the fluidics on our MoFlo are uncannily stable; who knows if we'd strike it rich again with a new sorter. You may also be asking, what's so great about XDP? Well, I'd never be able to explain with such elegance as Dan Fox could, so all I can say is track down the white paper Dan wrote, read it, then pick your lower jaw up off the floor. The big lure for me (besides the obsolescence of parts for the legacy MoFlo) was just the fact that we'd be able to operate with no/low hard aborts similar to the Aria, which, when paired with the higher number of droplets a jet-in-air can achieve, should allow us to sort faster and maintain high yields and purity. With our XDP upgrade, we also had all our PMTs changed, and threw on two new laser lines to boot. - Side note - We had one of these co-lase towers installed on our MoFlo, which is also a product of Propel Labs, that basically combines two laser lines so they can be run colinear into the 3-pinhold MoFlo setup. We chose to put on a UV and Red laser and run them colinear through the co-lase tower. This now gives us a 4-laser MoFlo (355, 488, 561, and 640) - End Side Note -
As far as the actual upgrade goes, the install went pretty smooth. It took 2-3 guys about 3 days to completely tear down the instrument to basically an empty table, and then install the PMTs, electronics, the touch-screen panel, and the computer. As with most installs/upgrades, we did have a couple hiccups, but they were taken care of immediately. I guess that's one good thing about working with a smaller company like Propel Labs. They can't afford to lose any business, so customer service is automatically very good.
We've been using the XDP now for about a week, and things have gone pretty well. We're still getting use to the touch-screen interface, and some of the new things in Summit, but overall, I'd say we made the right decision, and hopefully the MoFlo can dutifully give us another 10 years of service.
Once we've gotten into a rhythm on this thing and really test the bounds of speed, I'll post some data. But for now, enjoy a pic of the finished product below.
When I was contacted by the folks at Propel labs (who, evidently are a group of people from the original Cytomation company) that there was an upgrade path to the XDP electronics for the legacy MoFlo, I was thrilled. After about a year of begging for money from anyone that would listen to me, I finally secured the funding and was ready for the upgrade. So, why upgrade to XDP instead of buying a new sorter? Well, first of all, it was a financial thing. The cost of an upgrade is about 1/4th the cost of a new sorter. Secondly, the fluidics on our MoFlo are uncannily stable; who knows if we'd strike it rich again with a new sorter. You may also be asking, what's so great about XDP? Well, I'd never be able to explain with such elegance as Dan Fox could, so all I can say is track down the white paper Dan wrote, read it, then pick your lower jaw up off the floor. The big lure for me (besides the obsolescence of parts for the legacy MoFlo) was just the fact that we'd be able to operate with no/low hard aborts similar to the Aria, which, when paired with the higher number of droplets a jet-in-air can achieve, should allow us to sort faster and maintain high yields and purity. With our XDP upgrade, we also had all our PMTs changed, and threw on two new laser lines to boot. - Side note - We had one of these co-lase towers installed on our MoFlo, which is also a product of Propel Labs, that basically combines two laser lines so they can be run colinear into the 3-pinhold MoFlo setup. We chose to put on a UV and Red laser and run them colinear through the co-lase tower. This now gives us a 4-laser MoFlo (355, 488, 561, and 640) - End Side Note -
 |
| The remains of the MoFlo after the tear-down |
We've been using the XDP now for about a week, and things have gone pretty well. We're still getting use to the touch-screen interface, and some of the new things in Summit, but overall, I'd say we made the right decision, and hopefully the MoFlo can dutifully give us another 10 years of service.
Once we've gotten into a rhythm on this thing and really test the bounds of speed, I'll post some data. But for now, enjoy a pic of the finished product below.
 |
| The upgraded MoFlo XDP in all its polished glory! |
Thursday, September 23, 2010
Follow us on Facebook!!
Find the University of Chicago Flow Cytometry Core Facility group and feel free to leave any comments and questions you might have.
Wednesday, September 22, 2010
GLIIFCA Core Manager Meeting Preview - September 24, 2010
For those who will be at the meeting, I've put up my slides in PDF format here (sorry, I think the link works now) in case you wanted to follow-up with one of the tools we use in the core. For those who will not be able to attend, feel free to read through to see what types of tools we use at UCFlow to try to do more with less and be as efficient as possible. Check back here for updates from the Core Manager meeting and the rest of this year's GLIIFCA.
Thursday, September 16, 2010
Flow cytometry leads to everything!
The man on the right in this picture is Jeff Schneider. He used to be a technician here in the flow lab. Look at him now, playing with his band Darling at the Hideout tonight at 9PM. Rock on dude!
Flow cytometry leads to everything.
Subscribe to:
Posts (Atom)