UCFlow

What to do with aging equipment: Upgrade or Replace? And a Mini MoFlo XDP Review.

2012-01-10T15:23:00.000-06:00

I started writing this post specifically to follow through on a comment I made when talking about our upgrade of our aging MoFlo to the XDP platform (thanks for the reminder Carol). But then, I started thinking about all the equipment we've held onto and decided to upgrade and asked myself, was it really worth it? Before I answer that, let's lay out a bit of discussion on the matter, and then I'll finish up with my thoughts on our upgrade to the XDP.

When I think about equipment, I like to put things into 3 categories, namely Cutting Edge, Mid-Cycle, and End-of-life (EOL). I take all my equipment and shuffle them into these categories and move them around every so often as needed. This way, I can put things like service contracts, maintenance budgets, capital investment, and upgrades into perspective according to pre-determined criteria. For example, I'll stick instruments like our 4-laser Fortessa, or ImageStreamX into our Cutting Edge category. This means they probably won't require a huge maintenance budget since things aren't likely to break yet. However, they may need more personnel time because the applications performed on them are likely to be complex. I'll shuffle staff and training resources to those instruments. Mid-Cycle equipment are things like our 4 year old LSRIIs. Things are likely to start breaking and so they may eat up some service budget, but they are the workhorses and need to be running full-time. The applications are probably fairly routine, so they may not require as much custom tech time. Lastly, our EOL instruments are things like our ancient FACScan and FACSCantos, and it's these instruments on which we need to make decisions. Depending on the maintenance of these EOL'ed instruments, they may require varying amounts of service and since they may not be as desirable to use as the cutting edge cytometers, you'll need to determine how much money you're willing to invest to keep them limping along.

These EOL instruments can be a pretty decent consumer of budget and may or may not return all of their costs from recharge. It is with these instruments that we must decide; replace or upgrade (or I guess you could just let them die a slow death). You'll need to first determine if there is an upgrade path for your instrument. In the case of the MoFlo, this was a whole-hearted YES, thanks to the good folks at Propel Labs. Other instruments where this may be a possibility include FACScans and FACSCaliburs, which can be transformed into completely new instruments courtesy of Cytek Development. If there is not a path to upgrade, then the decision is an easy one. However, if you're looking into an upgrade, you'll need to weigh the costs against the benefits and definitely compare it to simply purchasing a brand new instrument. If you're going to shell out a bunch of money on an upgrade, it may make more sense to look into getting a new cytometer. Sure, you may have to settle with something a bit less powerful, but it'll be nice and shiny and (hopefully) problem-free for a few years.

So, let's put this all into practice with a retrospective look at our decision to upgrade our MoFlo. We had our MoFlo originally installed in 2000, and approaching 2008, it was definitely showing its age. Many of the buttons on the "rack" had fallen off, and it seemed like a waste of money to replace entire electronic bays on a rack to simply fix a button. In addition, parts to fix the MoFlo were somewhat scarce, and it looked like many of the components were approaching their demise. At the time, options for a new sorter were limited to the FACSAria, the inFlux (both from BD), the Reflection (from iCyt), and the MoFlo XDP (from Beckman Coulter). All of these instruments easily approached the $0.5Million mark, so buying a brand new sorter without an SIG or a generous donor was pretty much a long shot. Seeing as our MoFlo was still humming along just fine we decided to look into an upgrade. The goal going into this thought process was to have a sorter that would handle a lot of the cell line type sorts using GFP or other RFPs and perhaps a few phenotyping experiments. We did not have the expectation that it would rival our FACSAria and start performing multicolor phenotyping sorts as well as the Aria does. We also noted that our "GFP" sorts accounted for about 30% of all our sorts and guess what? We had 3 sorters; a perfect match. We were able to upgrade our MoFlo for about 1/3rd the cost of a new sorter and get a few more years of life out of it while we waited for the next big thing!

MoFlo-XDP Mini Review:

I can say that the XDP upgrade pretty much met our expectations. It handles most of our GFP/RFP sorts just fine, and is able to do a few more sorts on markers that are relatively bright. It by no means can resolve populations as cleanly as our Arias, but it does well enough for many things. The single best feature of the XDP is zero coincidence aborts. You may be thinking to yourself, well isn't the Aria marketed as having very low abort rates as well? It is, but when i say zero aborts, I really mean zero aborts, even when you have 30,000 - 40,000 events going through per second. The place this comes in handy are rare event sorts at high throughput rate. We can sort very rare populations and have a really good yield when compared to our Arias. What this really means, however, is that you have absolute control over your yield. If you need every single cell possible, you can run fast, have confidence that you'll be able to make a sort decision on every single cell, and using a yield sort mode, sort out every single cell. Sure, it won't be very pure, but at least you have them all and can decide to resort again if you need purity. Our specification for sorting yield is a 1% population with 70% yield using the purify sort mode (to achieve 98% purity or better). The max event rate able achieve this on the XDP is about 30,000 eps. The max rate able to achieve this on our Arias is slightly less (~22,000 eps). The touch screen is a bit annoying at first, but I've gotten use to it. The biggest problem with it is the implementation of the slider and up and down arrows. The slide is way too sensitive, and the up and down arrows are way too slow. This interface is used for adjusting things like frequency and amplitude and plate voltages. The new and "improved" Sort Master, dubbed Intellisort, works intermittently for us. It took a lot of playing around, but we can get it to hold onto a node pretty well these days, but for a while we completely ignored it. I still think this can be done way better, and apparently Intellisort II delivers, but I'm not going to hold my breath for that one.

So, am I happy with my decision? Absolutely! Would I do it again today? Not too sure. What it boils down to is, I spent a good chunk of change for a sorter that has 4 lasers and about 4 usable detectors at any given time. The need for the 4 lasers is pretty low, so I could get by with a 2-laser 4-color sorter and be able to do everything I'm currently doing on my 4-laser 10-color MoFlo-XDP. If I were given the option today, knowing what I planned to use the sorter for, I might check out the possibility of getting a brand new sorter that was stripped down to the basics for cell line transfection sorting. I'm thinking something like this perhaps might do the trick.

Is Compensation really necessary?

2011-12-19T15:54:00.000-06:00

For some reason, it seems like the idea of compensation gets so much 'publicity'. Everyone is always talking about compensation and how difficult it is. New users of flow cytometry tend to think of this idea as something so complex that they end up stumbling on this one idea before they even get started. So, let's get one thing straight right off the bat; compensation is easy. In fact, I'd say compensation is ridiculously easy today, now that you really don't have to do anything. You just identify your single stained controls, and your software package uses that information to compensate your samples for you. The real difficulty in performing flow cytometry assays is panel design - determining which colors to use and coming up with a panel where you have the optimal fluorochrome coupled to each antibody to give you the best resolution of your populations. In fact, I'd go so far as to say that in some cases, compensation isn't even necessary.

Wha, Wha, Wha, What??? That's right ladies and gents - compensation isn't even necessary (in some cases). And, I'm not just referring to the instances where you're using two colors that don't even overlap, I'm talking about straight-up FITC and PE off a 488nm laser. Now, before you stop reading and jump over to your Facebook feed let me just assure you that you first learned of the superfluous nature of compensation when you were about 5 years old. You see, analyzing flow cytometry data with or without compensation is nothing more than a simple "spot the difference" game you use to find in the back of the Highlights magazine while waiting to get your annual immunizations from the pediatrician. If you take a look at the figure below you may be able to recognize the left panel as the FMO (Fluorescence Minus One) control and the right panel as the sample. Spot the difference? Instead of seeing the sun missing on the left and then appearing on the right, let's just substitute a CD8-PE positive population for the sun. It doesn't really matter if the image is compensated, you're just comparing the differences between the two.

Let's make the comparison a bit more directly. Here we have some flow cytometry data showing CD3 FITC and CD8 PE. Our goal is to determine what percentage of the cells are CD3+CD8+. Obviously, there's some overlap in the emission of the FITC fluorescence into the PE channel when run on a standard 488nm laser system with typical filters. If I were to hand you this data set and pose the question of "What's the % double positive," you could employ the same strategy used above in the spot the difference cartoon without knowing a thing about compensation. The top two plots below are the FMO controls (in this case, stained with CD3 FITC, but not stained with anything in the PE channel), and the bottom plots are the fully stained sample. In addition, the left column of plots were compensated using the FlowJo Compensation Wizard, and the right column of plots are uncompensated. Were you able to "spot the difference"? If you take a look at the results, you'll see that either way we come up with the same answer. So what's the point of compensating?

As you can imagine, this is greatly simplifying the situation, and when you start adding more and more colors, you simply cannot create an n-dimensional plot that can easily be displayed on a two-dimensional screen. This could easily work for 2-color experiments - it could even work for 3-color experiments (maybe using a 3-D plot), but beyond that, you're going to have to do one of two things. 1. Bite the bullet and get on the compensation train, or 2. Abandon visual, subjective data display altogether and move to completely objective machine-driven data analysis. Compensation, much like display transformation is a visual aid used to help us make sense of our data, two parameters at a time. In our example above, we don't magically create more separation between the CD3+ CD8- and CD3+ CD8+ populations. The separation between them is the same, we're just visualizing that separation on the higher end of the log scale (when uncompensated) where things are compressed in one case, and on the lower end of the log scale (when compensated) where things spread. You didn't gain a thing.

10 Steps to a Successful Flow Cytometry Experiment

2011-12-12T23:00:00.000-06:00

I've been doing a good amount of application development recently and have had to "practice what I've preached." Those of us in the flow cytometry world, especially those in core facilities, like to pontificate all the do's and don'ts of flow cytometry, but how many of us have (recently) struggled through all the intricacies of perfecting a staining assay. I must say, I was a bit cavalier when I first agreed to set some protocols up for an investigator. The staining protocols weren't anything novel or difficult, it's just that I personally had not done some of the assays in quite a while. As I was going through the process I thought, hey, this is not as trivial as one might think...and I've been doing this for a loooooong time. I could only imagine what someone who is brand new to flow cytometry as a technique must feel like when their PI suggests they use this technology to investigate their hypothesis. So, I can put forth my top 10 steps to a successful flow experiment with some conviction, because I have now walked in your shoes.

I really wanted to make this a top-10, but as hard as I tried, I could only pare things down to 11. So, without further adieu I present to you;

10 11 Steps to a Successful Flow Cytometry Experiment

1. Read lots of protocols (not just the reagent manufacturer's protocol). Let's face it. If you ask a dozen people how to make a peanut butter and jelly sandwich, you'll end up with 12 different recipes. The same goes for FCM protocols. Everyone finds a different part of the protocol worthy of emphasis. If you read a few of them, you can start to put the entire story together.

2. Know which colors work best on your instrument. This is probably a bigger deal when you're using a core facility with a few different platforms. Let me tell you firsthand, no two cytometers are alike in their capabilities, not even two of the same model of cytometer. If you're lucky enough to have a flow cytometry core with knowledgable staff, make sure to ask them what their favorite 4, or 5, or 6-color panel is. They should also be able to tell you what the limitations of certain colors on a given instrument may be.

3. When designing your panel, look for max brightness with min spillover. Ok, let's say you know what sort of antibodies you want to run, and you know what's available, as far as hardware goes, at your institution. Now comes the fun part. You have a list of antibodies, and a list of fluorochromes - how do you match them up? You've probably heard the old adage, put your dim fluorochromes on the antibody that targets abundant antigen, and your bright fluorochromes on antibodies against sparse antigen. In addition to that you want to minimize spillover - fluorescence from probes that are excited by the same laser and whose emission overlaps. Spillover = Background, and Background = Diminished resolution. This takes some effort and a bit of know-how, so consult your friendly flow guru for help, or try out some of the new utilities designed to help with this process (namely CytoGenie from Woodside Logic or Fluorish from Treestar).

4. Titrate your reagents. What for? The manufacturer told me to use 5ul per test (usually 10^6 cells in 100ul of volume). Without jumping on the conspiracy theory bandwagon that reagent manufacturers tell you to use too much antibody so that you'll waste your antibody and have to buy more, I will say that I've found more times than not that the manufacturers suggested dilution is too concentrated. If you want to see why you should titrate your antibodies, check out the figure below. If you want to see how to titrate your antibodies, click on over to this prior entry to the UCFlow Blog.

CD4 staining of fixed human PBMCs at the properly

titrated concentration (Left) and the manufacturer's

recommended concentration (Right).

Example Staining Worklist

5. Outline your plan of attack. Make a detailed work list of your protocol. Generic protocols are good to help plan your experiment, but when it comes time to perform the steps of an assay, you really want a work list. As the name implies, this is a step-by-step recipe of how to execute the protocol. I usually include the step, duration, volume of reagent, temperature, etc... While you're performing your assay, take copious notes so you can fine-tune the protocol, adding more detail. The goal is to be able to hand this work list and the reagents to another user and they should have successful results. I like to do this in Excel and write in all the cell formulas so that I can type in how many samples I need to stain and have it automagically do all my dilutions for me. I also have a summary of the buffers needed and quantities at the bottom. See below as an example.

6. Always use a Dead Cell Marker. Dead cells can really screw up an analysis. I guarantee there is a color and assay compatible dead cell marker available for most every experiment you will do. There's no excuse not to use a dead cell marker, so please, please do it. It makes for a much nicer looking plot, and you really can't do good (dim) double positive enumeration without it.

Two-parameter plot without using an upstream dead cell

marker (Left) and the same plot after removing dead

cells (Right). Note the diagonal population extending

out of the negative population (encircled with a region

in the left plot)

7. Set up your FMO's as a separate experiment, not on your real samples. I won't discuss the merits of using an FMO control (Fluorescence Minus One), let's just assume you know that it's pretty much a necessity. What I will say is if you try and set up an FMO control on the day that you're using your precious sample, you're likely to either forget it, or omit it because you think you don't have enough cells. So, if possible, set up your FMO controls ahead of time on a different day so you can take your time getting everything set up properly. It'd be nice to include it every time, if you have enough sample.

8. Make compensation controls using beads. I'm a huge advocate of using capture beads to set up compensation. It's really a no brainer. I've written about this subject before. Even if your single stained controls look fine on cells, I'd still use beads because they're always consistent.

9. Acquire your samples nice and slow to achieve maximum resolution. If you go through the trouble of perfecting your staining procedure, now's not the time to screw things up. On a hydrodynamically focused instrument you'll want to concentrate your sample and run it slow in order to keep a narrow core stream and achieve optimal resolution. If you're using another type of flow cell (such as a capillary a la Millipore or an acoustically focused system like the Attune) you should be more focused on increases in background due to insufficient washing rather than a wide sample core.

10. Analyze your data a couple of different ways. Even if I have a clear idea of how to go about the analysis, I'm frequently surprised at how many times I've changed axes or started backwards and found I liked the new way better than the old way. Backgating is one way to help identify a rare population all the way up through its ancestry. Make sure to take advantage of your Live cell channel as well as gating out aggregates and removing any time slices where there may have been a drift in fluorescence.

11. QC your instrument and create an application specific QA protocol. Science is not about 1-shot deals. If it's not reproducible, it's not real. In order to give you the best possible chance of getting reproducible data you'll want to minimize the error contributed by the instrument. Quality control and Quality assurance cannot be emphasized enough. By doing something as simple as running beads at your application-specific voltage settings you can ensure that the instrument is in the same state as it was the last time you acquired these samples. For this, I typically use one of the peaks (peak 4, actually) of the 8-peak bead set. After I have the samples acquired with the proper voltage settings, I run the beads, create target channels for the peaks and save it as a template. Next time, all I need to do is dial in the voltage to put the beads in the target. You'll also want to make an Acquisition template and probably an analysis template too.

Well, there you have it. Hopefully this will help you focus your attention on some key aspects of setting up a well-thought-out flow cytometry staining protocol. Of course, this merely scratches the surface of all the things you need to think about. Did I miss something major? Feel free to leave a comment with your #12, #13, and beyond.

The Year of Acquisitions

2011-12-02T10:31:00.001-06:00

Like most industries, the Flow Cytometry Industry appears to be shrinking, in that the number of players on the industry side of things is getting smaller. For many years, there were a few big players, namely Becton Dickinson and Beckman Coulter (who they themselves were products of mergers - BD+Ortho, and Beckman+Coulter). They made instruments and reagents and pretty much sold the whole package. Seeing the potential for others to capture some of the market share, we experienced a growth of smaller start-ups, either focusing on the hardware or the reagents. Companies like Cytomation (maker of the MoFlo), and Guava on the instrument side of things introduced some nice products and created some much needed buzz. A major impact of these companies was that it forced the major companies to invest in R&D and come out with more competitive products. On the antibody side of things, reagent-focused companies like eBioscience and Biolegend gained popularity. But, I think a real turning point happened when little-known Accuri Cytometers exploded on the scene with a low-cost, small footprint cytometer with capabilities similar to a FACSCalibur. They took a page from the Guava playbook and targeted individual labs instead of the typical cytometer purchaser - a core facility. Soon other companies were seeing the success of these platforms, and the much larger market outside of the core facility. Companies like Stratedigm, Life Technologies and iCyt started offering smaller sized, less expensive cytometers. It seemed like the cytometery industry - both on the instrument and reagent side - was expanding. This lead to competition and innovation. The old standby's like BD and Beckman Coulter were forced to come up with new and exciting products to maintain their market share. And then the recession hit.

So, what happens in a recession. Well, contrary to what you might think, many companies do just fine in a recession. Of course their growth may slow, but then they also tend to accumulate capital as well. In fact many companies wind up in a situation where they have lots of cash on hand and are sort of waiting to see what's going to happen. John Waggoner explains in a USA Today piece (http://www.usatoday.com/money/perfi/columnist/waggon/2011-05-05-cash-in-on-mergers-and-aquisitions_n.htm) that this past summer, it was estimated that companies in the S&P 500 stock index had a combined $940 Billion in cash. I postulate that the well-established cytometry companies were/are in a similar boat...but to a much lower degree.

Mr. Waggoner goes on to explain, companies with cash-on-hand basically have three things they're going to do with it.

1. They can reinvest in the company, hire more people, build more plants, funnel it into R&D, etc... However, with funding becoming more and more scarce, there's not enough demand in the market to warrant such reinvestment.

2. They can return money to their investors in the form of dividends. Some companies are doing this, but probably in moderation.

3. They can buy another company to position themselves for the recovery. Mergers and Acquisitions are a pretty huge business in recent years. In total, M&As are running at a $1.6 Trillion pace for 2011. A good chunk of this is happening in the healthcare sector.

Bingo. Herein lies the recent increase in mergers and acquisitions. For example, Accuri raises ~$30 Million to get their business going, BD sees the threat and buys them for $205 Million (not a bad ROI for the Accuri Investors). BD removes the threat, and clears the way for its new flagship, small footprint, easy-to-use cytometer, the FACSVerse. This works for reagent companies too. Affymetrix buys eBioscience, EMD-Millipore buys Guava and now Amnis, Life Technologies licenses the acoustic focusing technology to build the Attune, and on and on it goes. Even bigger name companies like Sony and Danaher are getting into the game. Sony purchased iCyt to see if it can get its foot into the biomedical research arena, and Danaher purchased Beckman Coulter for who knows what reason. At any rate, it seems like the industry is attempting to go back to the old days where you'd do all your shopping at one company. Buy your instrument, reagents, analysis software, and all the rest from one company. You'll end up having BD labs, Millipore Labs, Life Technology Labs and maybe even Beckman Coulter Labs. A necessity in the current environment, but I'm sure things will oscillate back to the innovative start-ups taking on the big-boys once again. So, who's next to be gobbled up? I'm sure companies like Stratedigm, Blue Ocean, and Cyntellect are hoping their phones will start ringing.

Options for Flow Cytometry Training - FloCyte Review

2011-11-21T19:41:00.001-06:00

Flow Cytometry (FCM) isn't the easiest technique to learn. It actually takes quite a while to master both the hardware and software components to sample acquisition and data analysis - let alone the applications utilizing the aforementioned instrumentation. For many users of flow (in an academic setting) their first encounter with FCM is likely through a core facility, whereby they'll receive some instruction on how to operate an instrument and then how to analyze the data they collected. The type and quality of this training varies greatly. Some institutions I'm familiar with have multi-day courses with wet lab sessions and hands-on instrument time, while others attempt to provide a theoretical base and then do a bit of hand-holding for a few sessions. The success a user may achieve greatly depends on his or her resourcefulness and overall aptitude for technology. Some people pick it up quickly; others struggle for years. I will say that training users in a busy core facility is a huge drain of time and resources. In our core, for example we basically have an entire F.T.E. just providing training and consultation, so I'm sure that in smaller cores, where it's just one or two people, training has to be an even greater burden. The question then becomes, how are we to provide the necessary training and attention our users require with the limited time and personnel resources characteristic of a core facility?

There aren't too many options. Before I jump into an assessment of the FloCyte courses (which is the whole point of this post) let me briefly highlight other possibilities. FYI, I've personally attended all 3 types of training sessions and have viewed all the resources in #4.

1. The Annual Course in Flow Cytometry - This weeklong course alternates between Los Alamos National Labs (or the University of New Mexico) and Bowdoin College in Brunswick, ME. It is really geared towards users of the technology who already have a basic understanding of the technology. Also, it focuses on the applications of flow cytometry rather than operation of a flow cytometer, however numerous sections also delve into the hardware components. There's a pretty cool lab where you can assemble your very own (fairly crude) cytometer. The cost of the course is about $1800, which includes dorm-style accommodations and meals (transportation is not included).

2. Vendor-specific instrument/software training - Most vendors will provide training for their hardware and associated software. When you purchase an instrument, you might get some free training included with the purchase, but additional training is going to cost you. As you'd expect, the training is geared towards the operation of that vendor's hardware. If you were using multiple cytometers from different vendors, this obviously wouldn't be ideal, but if you were using a single platform it might be a good option. The vendor training will also include some of the basics of cytometry, but again, it will be skewed towards their instruments, their reagents, and their idea of the technology. It's also pretty expensive, sometimes as much as $2500 per person.

3. Training courses at meeting - Typically when you go to some of the bigger conferences they'll have some workshops on FCM. Certainly at the CYTO meetings you'll have the opportunity to attend training sessions on various topics. Also, some of the immunology focused scientific meetings will have some FCM training associated with them (for example, the AIC meeting in Chicago). Cost for this training is variable, however it's usually limited to conference attendees, so unless you were already planning to attend the conference, it might be really expensive.

4. Online utilities - There is quite a lot of information freely available on the web. You can certainly start at the Purdue University Cytometry Laboratory web site, where there are a bunch of powerpoint slides, movies, and resources freely available. In addition, companies such as Becton Dickinson, Life Technologies, and Beckman Coulter offer overviews of flow cytometry and flow cytometer technology. Note that the above links are linked directly to the company's training/support page with the intended materials. Although these online utilities are readily available and free, you lose the benefit of asking questions and interacting with people who can tailor the training to your specific needs.

So now, I'll walk you through my experience with the FloCyte Training course offered by FloCyte Services. I attended the Comprehensive Training Course from 11/15/11 - 11/17/11 held at Spherotech, Inc. I won't bother taking up space here to give you the rundown of the company and the mission of the training courses. You can read all about it here. However, I will note that I attended the Comprehensive training course, which is designed for novice users of flow. You can see the course curriculum here.

Day 1, as you'd expect, goes over the basic components of flow cytometry. This is done is a pretty common fashion, and anyone who's gone through the powerpoint slides on the Purdue University Cytometry Laboratory web site will recognize the format. 4-components, Fluidics, Optics, Electronics, and Data Analysis. All the standard material you'd expect to be here is here. There was however at least one pretty critical omission - multi-laser systems, laser delays, and how fluorescence emission is spatially separated. I know this was briefly mentioned during one of the sections, but there was no figure, no reiteration of how it's possible to look at two colors with the exact same emission simultaneously because they're excited by spatially separated laser beams (e.g. PECy7 and APCCy7). When we broke into small groups to take a look at some of the hardware, I spent most of the time explaining to my other group members how this works. They were very confused. The graphics used to talk about emission filtering where all systems like a FACScan or FACSCalibur, which don't have spatially separated beams, and all the light goes through the same "pinhole". Also on day 1, we finished up with a mathematical explanation of compensation, which went horribly wrong. The math is complicated and it's probably not something basic users need to understand in order to compensate their data correctly (or, should I say, let FlowJo compensate their data correctly). Lastly, there was no mention of 1 very critical component to flow cytometry, Quality Assurance and Quality Control. In all, the basics were handled just fine. I will say, though, that it seemed to move pretty slow. I think for the amount of information covered in that first day, it could've have been condensed into a half day. For example, I feel like the flow basics class given at UCFlow is comparable in it's scope but is completed in about 1.5 -2 hours.

Day 2 brought in a plethora of applications and tried to reinforce some of the concepts from day 1 while explaining how those concepts effect how you think about the applications. I think this way of presenting the information is really good. When we're talking about immunophenotyping, we're also talking about compensation, background due to fluorescence overlap, non-specific binding, etc... When we're talking about cell cycle, we're also looking at doublet discrimination, coincidence, sample core size, etc... Here we also start tackling the necessity of controls, including comp controls and the always popular FMO controls. My big issues with this section solely revolved around the figures. Many of the figures were at best poor representations of the idea being put forth and at worst blatantly misleading. This was especially noteworthy in regards to an explanation of biexponential display transformation. In another instance, the instructors were driving home the idea of how we are to never use quadrants to perform gating on our plots and the very next slide describing FMO controls was filled with quadrants used as gating. A bit contradictory.

Day 3 was all about stats and panel design. The stats part was very straight-forward and pretty easy to follow. The panel design section was good, and covered many of the issues that arise when trying to put together a multicolor panel. There was an introduction to a utility from Treestar called Fluorish (which I'm not going to complain about because I like it) however there wasn't any real mention or demonstration of other available utilities like Chromocyte and CytoGenie. Also, we spent some time going through some data analysis strategies using FlowJo.

The cost for the 3-day Comprehensive course is $700. The beauty of the course is that it's brought to you (either your institution can host it, or it is hosted nearby) so you don't have to factor in airfare or hotel costs. But, you'll have to remember that you're getting a comprehensive theoretical overview of flow cytometry, you are not learning how to operate your specific cytometer. So, if you didn't have a core facility around to show you how to open up FACSDiVa and adjust voltages on your LSRII, you'd still be pretty clueless on how to run your first FCM experiment. Another positive about the training is that it is modular such that you can attend just days 1 and 2, or just 2 and 3, or even just day 3. That way if you have some basic knowledge already, you can skip day 1 and just attend days 2 and 3. Lastly, I'll mention that there are a bunch of other, more advanced courses available outside the comprehensive course, including a multicolor compensation course, a course on "phosflow" assays, and even clinical flow cytometry.

The instructors are well-respected flow cytometry professionals with years of experience under their belts. They presented most of the material in a clear and concise way. There was, at times, some confusion regarding what a figure was trying to describe, but this was due to the fact that the slides were recently re-done and the instructors were not 100% comfortable with them. I feel like I want to give them a pass on that, but then again, I did pay $700 on this course and expected a very polished delivery. All things considered, they did an excellent job.

I could see this working in a couple of ways. 1. You get some initial training on how to operate your cytometer from your core facility and then attend days 2 and 3 of the comprehensive course. 2. You could attend the entire comprehensive course and then go through the specific instrument training given by your core facility. 3. Get trained by your core, start running experiments, and then jump in on one of the advanced courses offered by FloCyte. If you're not fortunate enough to have the support of a core facility, then this makes the FloCyte courses even more attractive. Relying on them for the basic theoretical training, and then the instrument vendor for training on the actual equipment is probably your best bet.

Cytometer Service Contract or Self Insure: the Wal-Mart effect.

2011-11-11T14:56:00.001-06:00

Instrument maintenance and repair is typically not a huge factor when deciding on a piece of equipment to purchase. People are much more concerned with the practical things like how many lasers can I put on, how fast can I run my samples, or more simply, can it handle the applications I plan to run? Even after we have the instrument installed in the lab we're not really thinking about maintenance and repair because we're on the "full-warranty high." If something breaks, what does it matter? The company will come out the next day and repair it at no cost. Right about the halfway point through the warranty period the thought hits you - I'm going to have to start paying for service on this thing. Herein lies the dilemma.

Although there are many variations, in general there are two schools of thought here. The first involves some level of service agreement (full, partial, lasers only, instrument minus lasers, etc...) and the second is akin to an "insurance" plan. By the way, before I go on, I should state that I'm writing this from the standpoint of a private academic institution (namely the University of Chicago), however private companies, public institutions, or individuals may have a vastly different experience. Let me briefly explain these two systems of instrument maintenance.

Service agreements. About 6-8 months into your warranty period, a friendly company representative will contact you to try and sell you on a full service contract. This basically extends the type of service experienced during the warranty period. Labor and parts will be covered under the service contract costs you pay annually. Be sure to get a list of what are typically called 'consumable parts'. These items are parts that will not be covered under the service contract. Consumables are commodities that are intended to be used up quickly and therefore are not parts that could undergo some type of failure. It is this failure of a part that is covered by the service contract. Consumables can be expensive; sometimes as much as $1000 - $2000 for a single item that may only last 6-12 months. You'll need to be sure to add these costs to your total cost of ownership. Full service contracts are fantastic. You get rapid response times, an endless supply of new parts, and generally I find the quality of service is of a higher standard. The downside is the expense. You can plan on spending about 10% of the original purchase price yearly on a full service contract, which means that after 10 years, you'll have bought the instrument twice. You can also look into service contracts that cover only parts of the instrument, such as a 'lasers-only' contract. This may cover some of the major expenses that might hit, but some of the routine fluidics issues or electronics issues would still need to be paid out-of-pocket. Lastly, you don't need to rely solely on the Original Equipment Manufacturer (OEM) for service. In some cases, third party companies will either provide the service agreement (serve as a middle man between you and the OEM) or there are companies that can actually come out and fix some of your older generation instruments.

Insurance. If you pass on the service agreement route, either with the OEM or a 3rd party company, you'll need to carefully make a plan on how you will pay for problems that pop up. This can be done by including a line item on your budget and simply inserting the cost of the service contract. Then you'd need to pay for any repairs using those available funds. If you don't use all the funds then you have a surplus and possibly a way to do some upgrades or save it for a rainy day. If, however, you end up paying out-of-pocket more than you have put away as insurance then you could have some trouble with your institution. The insurance method also has some unintended consequences including the possibility that your service calls may be bumped to the bottom of the list if the OEM services customers on service contract first. Secondly, I've noticed that the field service engineers tend to do the minimum to get the instrument functional again. This is not to say they're lazy or anything, they're actually doing you a favor by not replacing non-essential parts, and performing the work quickly so the hourly labor charge is not too high. However, this sometimes leads to more frequent trips to a site to fix a related part that breaks shortly after the instrument was put back into service.

So, what do we do at UCFlow? Well, a hybrid, of course. If you can anticipate which instruments will likely have more problems over the years, then you can keep your instruments running for many years without hassel for a lot less money. Seems impossible, but here are a few tricks. Obviously, the first thing you're going to do is monitor performance very carefully during the warranty period. If odd things are happening monthly, or even quarterly, it may be a good idea to consider a service contract. If you can find out from current owners of the same model instrument whether they have many service calls, that might help make the decision. Also, if you or your lab is familiar with the innards of the cytometer and aren't afraid to do things like replace valves, regulators, or even lasers then you should be less likely to buy a service contract. Lastly, the more instruments you have, the more money you'll be wasting on service contracts. Let's say you have 6 instruments, and the service contract is $15,000 each ($90K total). It's unlikely that all 6 cytometers will have multiple issues in a given year, so let's say you have 2 instruments with major problems (multiple service calls with big ticket items totaling $30K). The other 4 run pretty smoothly, and maybe require another couple of service calls for minor issues ($15K). If you pay out-of-pocket then you'll basically be paying 50% of the cost of a full service contract. This might be a good year; some other years might not be so favorable. However, it's likely that many years you'll be under budget and a couple of years you might be over budget.

As an example, I'll share a few stories of my experience. We had multiple 1st generation FACSCantos that were breaking down monthly. We were actually paying more out-of-pocket than the cost of a full service contract, so we went ahead and put them on contract. This was a no brainer. We also had an old LSRII that, over the course of 6 years had not had a single service call placed on it. All we've had to do is perform the standard Preventative Maintenance (PM). We never had a contract on this instrument. After 6 years of spending nothing on this instrument, we had 2 lasers die at the same time, which required replacement at the cost of $50K. The service contract cost was $22K per year, so 6 years times $22K = $132K, and actual costs were $50K, a 62% savings. It is a situation like this that tells us to err on the side of NOT getting a service contract until an instrument proves to be unreliable. Once it is deemed unreliable we either place it under service contract, or get rid of it and find a more reliable alternative. It sometimes seems like a gamble, and if I only had 1 or 2 instruments, I'd likely have them on service contracts, but since I have the luxury of duplicate technology and the power of numbers, I'm able to take that gamble and the odds are usually in my favor.

By the way, of the 16 instruments we have in the lab, 3 are on service contract (only the aforementioned early generation FACSCanto-A). We're able to save money by having a high number of instruments. We can also negotiate better contracts if desired. Larger volumes typically lead to better prices per unit. This is what we call the Wal-Mart effect. If that's not your case, then you'll likely want to lean more towards the service contract route.

The most sensitive Cytometer available?

2011-10-25T21:34:00.000-05:00

Recently, we've had a pretty good bump in the usage of our ImageStream X (Amnis, now a part of EMD-Millipore), but many of the new users are using the technology to confirm things they're seeing on the conventional flow cytometers. So, needless to say, I've been doing a bit more phenotyping on the ISX instead of the usual nuclear translocation or apoptosis assays that we typically do. In doing so, I was reminded of some comments thrown out by Amnis at the 2011 CYTO meeting saying (and I'll paraphrase) the ISX, and by extension the FlowSight, is the most sensitive cytometer available. The evidence of such a claim was a screen grab of good ole 8-peak beads (Please don't get me started). So, I had some data that I recently collected and thought I'd try and validate those statements with some data that makes sense to me.

It's a really simple example, but in short it involves a surface marker (coupled to PE), a Live/Dead dye (Green) and a Nuclear dye (Violet). By conventional flow cytometry, the PE signal was pretty weak and the user was skeptical that the staining was "real." So, the idea was to make sure the cells were live (Green low/neg), were actually cells (Violet pos) and had surface staining of PE. After going through the normal groups of gating, it came time to look at the PE signal. Surprisingly, it wasn't bad at all (especially with the 561nm laser cranked up to 200mW), however there were some dimmer PE+ cells that were hanging out a bit too close to the negative.

I remember having a discussion with other people in the lab about using carefully calculated masks to pull out the membrane staining and completely removing the cytoplasmic and/or nuclear background which should bring the negative population pretty much down to zero while retaining the specific PE positive fluorescence. This procedure is actually pretty simple so I'll briefly explain it, and if this whole concept of image masking is foreign to you, just think of masks as parts of a cell defined by morphology within which you're going to measure fluorescence. This is very different from flow cytometry where you can only measure fluorescence from the entire cell regardless of where that fluorescence is coming from. With this data, I'm creating a membrane mask, which basically looks like a ring encompassing the outside of the cell. This should retain most of the specific PE fluorescence and remove both background from intracellular autofluorescence, but also background from the nuclear dye and live/dead dye. The figures below demonstrate these findings.

The figure to the left is the originally analyzed data. On the far left is just a gallery of images that show the different fluorescence (the green live/dead wasn't show since dead cells were gated out). The top dot plot is a simple SSC/PE scatter plot to show the distribution of the negative and positives. The image just below is showing the mask used (bluish semitransparent shape overlaying the PE image). Below that is the ungated population showing the Live/Dead Green fluorescence spilling into the PE channel using the whole cell mask. And lastly, a histogram showing the PE Fluorescence. Altogether, a pretty straightforward analysis. However, I wanted to see what would happen if I restricted the PE mask to only the membrane area, so that is what is shown in the figure below. Now, it's important to note that this is the exact same data file, analyzing the exact same group of cells. The only thing changed here is the mask on PE, which is now shown as a ring overlaying the membrane of the cell. If you now look at the SSC/PE scatter plot at the top, you can see the dramatic tightening of the negative population, which implies a reduction of the high autofluorescence cells that were trailing to the right of the negative population in the total cell mask. Another benefit of this restrictive masking strategy was the reduction in the spillover of the green dye into the PE channel as shown in the ungated Live/Dead Green versus PE plot. And lastly, when you look at the histogram, you can see unequivocally the increase in separation between the negatives and positives. To drive the point home a bit more, we can overlay the two histograms so you can see exactly how they match up. Notice that there is a reduction in the intensity of the positive population as well, but this is likely a similar reduction in background fluorescence as is seen in the negative population. The key here and really in all of flow cytometry is RESOLUTION. This is, in fact, what most people are really thinking about when they say 'sensitivity.'

So, can we confirm the original statement here about these imaging cytometers being the most sensitive cytometers available? Well, I'm not sure I'm ready to crown this instrument as the winner just yet, but at least in some circumstances, the ability to only analyze the part of the cell that is actually stained or not stained could prove to be an extremely vital tool especially if you need to resolve dimly stained cells from unstained cells.

Counting Cells with the EMD-Millipore Scepter 2.0

2011-10-13T16:21:00.004-05:00

I recently had the chance to play around with the Scepter 2.0 Automatic Cell Counter from EMD-Millipore. The Scepter uses the Coulter Volume principle to count cells in a microfluidic chamber connected to a handheld device. I'm basically using it for things like confirming pre- and post-sort cell counts, as well as counting cells being passaged and primary cells such as PBMCs and splenocytes. The device itself is basically shaped like a pipetteman, and even has a plunger type action which simulates pipetting.

EMD-Millipore Scepter 2.0

To use the device, you need to attach a single-use 40um or 60um sensor, which provides the microfluidic channel through which the sample is passed. Once attached, you simply hold down the plunger, submerge the sensor tip into a sample volume of ~100ul and then let go of the plunger. It takes up about 50ul of your sample through an orifice in the sensor and measures the volume of the cells. It then plots the volume (or diameter) of the cells in a frequency histogram displayed right there on the device's built-in display. Using a click-wheel on the finger grip side of the scepter, you can adjust the low and high bounds of the histogram in order to remove small (dead/debris) and large (aggregate/larger cells) events. Once you set these bounds, it displays the event number/mL at the bottom of the display. The sensor tips are single use and each have a range of cell sizes and sample densities it can handle. The 40um tip is geared towards cells with a diameter of 3um to 17um and a cell density of 50,000 cells - 1.5x10^6 cells per mL. The 60um tip can handle cells with a diameter of 6um to 36um and a cell density of 10,000 - 500,000 cells per mL. The handheld unit can store up to 42 histograms, but this data can be downloaded to a computer and analyzed with the Scepter Software Pro (Mac/Win - Free!). In the desktop software, you can add info like original volume, dilution factor, sample names etc.. You can also re-gate and overlay histograms to create handy figures. When you have everything set up, you can export reports in table format which you can open up with Excel or other spreadsheet programs.

Scepter Software Pro Screenshot

So, how does it work? Well, it certainly counts things very accurately. Previously, I've found that my MoFlo XDP reports sorted cells really well (at least when the side streams are behaving themselves), and I have lots of data comparing sorter counts to counts using a standard coulter counter or even counting cells on a conventional cytometer using absolute counting beads (which I get from Spherotech, by the way). The only drawback is you don't get a live/dead report like you might with visual-based systems (e.g. Hemacytometer counts with Trypan Blue, Coulter's Vi-Cel, Invitrogen's Countess, or even Nexcelom's Cellometer). Sure, you can sort of approximate what's live and dead using volume or diameter as a discriminator, but all the profiles I have been collecting don't really show a clear distinction. It's certainly not like staining some cells with PI and throwing them on a FACScan with counting beads. But, with that said, the system worked pretty darn well. Getting back to the accuracy thing, it matched my counts from a sorted fraction on my MoFlo to within 1%. That makes me feel good in two ways: 1. My MoFlo is sorting well, and 2. The Scepter can actually count really well in a very short amount of time (30 seconds by the way). I did have one small snafu (described below) which was giving me some really weird results, but outside of that, it performed as advertised. The one thing that I might have to complain about is the cost of the single-use, disposable sensors. $3 a piece. Ouch!

Blue histogram represents a collection in the upright

position, which leads to low end bubbles showing

up and screwing up the counts. The green histogram

repesents a collection up-side-down and no bubbles

ending up in the chamber leading to way more accurate

counts.

Now, about that snafu. What I kept finding was after the sample was loaded into the sensor, and then the sample started traveling through the sensor orifice into the counting micro-channel, I kept seeing bubbles creep in there. The effect of this was I'd start getting these really low volume events piling up near the end of the counting process. It was a small number of low volume events that I could probably gate out (see figure below), but it still messed things up for me. Since I was doing a 1:10 dilution (10ul sample, 90ul buffer), when I back calculate (or better yet, let Scepter Software Pro back-calculate for me) the concentrations, I was off as much as 1x10^6 cells (or a 12% swing in total cell counts). To solve this problem, I made one modification to the collection process. As soon as the sample was loaded into the sensor (it beeps at this point), I immediately flipped the entire Scepter apparatus upside-down as to force any air that begins to enter the sensor to remain near the tip and not enter the orifice and microfluidic channel. This got rid of all the air bubbles and my counts became extremely accurate. In one case, my MoFlo told me there should be 8.02x10^6 cells, and the Scepter counted 8.01x10^6 cells. This made me happy. To see this awesome flip move in action, check out the video below. I apologize for the sound, I was filming this in my sorter room, which has the gentle hum of a twin diesel engine for background noise. Also, you'll just have to trust me when I say "see the bubbles." UPDATE: After playing around with volumes a bit more, it's pretty evident that you definitely need 100+ microliters of volume in your tube. I could get bubbles every time if I only had the requisite 50ul of sample, but if I had 100-120ul, I almost never got bubbles. With this volume, there's no need to turn the scepter upside-down.

So, in all, I think this product was successful for what my purposes were. It's small. The counting process is fast. I can offload the data to my computer, and the counting was very accurate (as long as I remembered to hold it up-side-down to avoid the bubbles). Will I continue to use it? I guess it sort of depends on whether or not I can get over not 'knowing' the %live/dead. For what I'm doing, that's probably fine, but could another option be just as easy and accurate and cheap AND give me live/dead? To be determined. I will say that I've used early versions of the Countess and the Nexcelom, and neither impressed me so much as to make me want to buy one immediately. Hopefully I'll be able to check them out again and perhaps put together a head-to-head review.

GLIIFCA 20 Wrap-up.

2011-10-05T14:11:00.001-05:00

If you're unfamiliar with the Great Lakes International Imaging and Flow Cytometry Association (GLIIFCA) meeting, you can check out this year's program online here. It's sort of a morph between a technology focused user group meeting and a smaller scale scientific meeting. The focus really is on the utilization of our technology (which I'll refer to under the umbrella term Cytometry) in clinical, translational, and basic research. There is also a strong cytometry vendor presence; about 30 different companies bringing their latest and greatest products. If you'd like to see who attends and supports the association, you can see a list of sponsors on the GLIIFCA site. A part of the meeting that's always a bit disconcerting for me is the Friday night Industrial Science Symposium, which is code-language for "vendor sales pitches." It's been pretty poor some years and not-so-bad others. It really depends on the presentation and the quality of information put forth. You can tell some people are up there literally just trying to sell a product. A good presenter will educate the audience so that the individuals sitting in the chairs come to the conclusion on their own that this is the product they need. And I have to say, we witnessed one of the best examples of this last Friday night in a presentation given by a Chicago-favorite, Kelly Lundsten from BioLegend. Great talk, and actually a pretty good session in total.

A Slide grabbed from Janet Siebert's
(Cytoanalytics) Presentation at GLIIFCA 20

The "theme" of the meeting was Cytoinformatics (as opposed to Bioinformatics). As far as the scientific program, it was the first time I found myself thinking, maybe these informatics people aren't wacked. I hear what they're saying, but it usually doesn't strike a chord with me. The basic idea is that you're generating tons of data of various kinds that needs to be quickly integrated in a consistent format in order to support analysis and subsequent decision-making. And I think my resistance has always been in the format of, "Well I don't really generate THAT much data, so I don't have to worry about this stuff." After sitting through a few examples of data generation from some groups that I know pretty well, it got me thinking. The quantity of data can be pretty big even if you're only doing 8-12 parameter flow cytometry or less. This isn't something only for the 18-parameter groups, it's for everyone. Besides the flow data, it would be nice to integrate this info with subject info, imaging info, genomics info, etc.. I think what was pretty successful for this meeting is the fact that it was setup in such a way that you could see the progression of ideas surrounding management of data. 1. Here's the problem: People collect lots of heterogeneous data types. 2. Here's the types of tools needed: Data warehousing, including dimensional models, ETL (extract, transform and load data), and end-user tools to read the relational database. 3. Here are some examples of how people are using these tools with real data and how it impacts decision-making. That was basically GLIIFCA 20, Symposium 1, 2, 3. Kudos to the program committee.

UCFlow's GLIIFCA 20 Poster

There were also a pretty good crop of posters presented this year, including mine (which won a poster award, thank you very much). Two of them which stuck with me were the "Increased number of laser lines on your cytometer might mess stuff up, so be careful" poster and "Look at this awesome temperature control/antagonist injection apparatus I soldered together with some parts from Home Depot" poster. I'm paraphrasing the titles, of course, and you can find the full poster abstract in the GLIIFCA 20 program linked above. The first one is from the folks just up the road at Northwestern (Geoff Kraker and James Marvin), and the second one comes to us from Roswell Park courtesy of Ed Podniesinski and Paul Wallace. The UCFlow poster was about how "I can't stand looking at QC data, so I'll start using cool Google tools and graphics to make it more interesting and maybe I'll stick with it longer."

So, there you have it. Another year, another GLIIFCA. For the record, this was my 11th GLIIFCA attendance. I have officially attended a majority of GLIIFCA meetings.

Safety; It's not to be taken lightly

2011-09-22T19:10:00.001-05:00

I'll begin by saying, I definitely need to pay closer attention to the various safety concerns in a lab. All too often we sacrifice our own safety in order to get things done quicker; cutting corners, thinking I'll be careful. And then, bam, you have an incident that you regret. Fortunately, I haven't had to deal with this first hand, but what I'm going to describe here happened close enough to home that it caused me to pause for a minute and evaluate my own techniques and protocols in the lab.

Perhaps some of you are aware of a recent incident at the University of Chicago, where a scientist became infected with the same strain of bacteria that is being studied in the lab (B. cereus). According to information published on the Science Magazine site, the infected individual was not even working with the microbe but may have transfered it to an uncovered wound via a spill (http://news.sciencemag.org/scienceinsider/2011/09/university-of-chicago-microbiologist.html). I believe the infected person is going to be fine and needed to undergo surgery to remove the infected tissue, so that's positive. As a result of this (and another incident just 2 years ago), the PI is moving these sets of experiments to Argonne National Labs in the Howard T. Ricketts lab, where they are also running experiments on Plague, MRSA, and Anthrax.

It was roughly two years ago to the day that a researcher in this same Laboratory at the University of Chicago died from exposure to an attenuated form of Y. pestis. In this case, the researcher may have felt a bit safer than he was, since the strain was determined to be non-lethal. His co-workers admitted that his glove wearing practices were inconsistent at best. It just so happened that he also had an undetected/untreated condition known as hemochromatosis, or an overload of iron in the blood. It may have been this overload of iron that allowed the attenuated version of the bacteria to become virulent (http://en.wikipedia.org/wiki/Malcolm_Casadaban).

So, as you can see, we have plenty of examples of the potential threat to our safety and those around us, and we should use examples like these, not to place blame on those who made mistakes, but to remind us of the importance to slow down and think about what we're doing and what we need to do to stay safe. There's really just two reasons why incidents like this happen; Carelessness or Ignorance. You have to be aware of what you're working with. Ask questions if you're unsure. Educate yourself. Nothing is so important that you cannot take the extra steps to make sure you and those around you are protected as much as possible.

Those who work in your safety office are not out to get you. They're here to educate first and foremost, and yes, to enforce standard operating procedures for your protection. In perusing our own safety department's web site, I stumbled upon this - Shared Responsibility.

Mission: http://safety.uchicago.edu/about/index.shtml

Environmental Health and Safety provides services and support for efficient, effective, and compliant work practices, while promoting a culture of shared responsibility by students, faculty, staff and visitors for a healthy, safe, and environmentally sound educational and research community at the University of Chicago.

And specifically regarding Laboratory Safety, they have this to add: http://safety.uchicago.edu/labpersonnel/index.shtml

Research is one of the two main missions of the University, the other being education. Lab personnel are integral in the creation and maintenance of a safe laboratory environment. They are responsible for ensuring safety in laboratories and lab support areas on campus and within the Medical Center. This responsibility includes:

Being familiar with University emergency procedures;
Responding appropriately in the event of an emergency;
Being familiar with Environmental Health and Safety policies and procedures;
Maintaining a safe laboratory environment;
Knowing the hazards of the materials and/or equipment being used;
Following all safety procedures in the laboratory environment;
Selecting, using and understanding the limitations of personal protective equipment;
Reporting any unsafe conditions to your supervisor and/or Environmental Health and Safety; and
Reporting any job related injuries or illnesses to your supervisor or Human Resources Administrator immediately; and
Participating in all required safety training.

So, hopefully if you've taken the time to read through this, you can certainly take the time to re-evaluate the procedures in your lab. Make a plan to educated your staff and those around you, and open the lines of communication between your lab and those who have been tasked with the safety of your institution. Stay safe!

Where's my Dream Cytometer?

2011-08-10T12:41:00.000-05:00

Have the market research groups recently been clamoring at your door? It seems like a weekly request via email or phone call to take "10 minutes" to answer some questions about "the future directions of flow cytometers and associated reagents." I've answered these calls so many times in the past few months that I'm starting to sound like a broken record. My hope is to perhaps just send them this link instead of spending time scoring questions on a scale of 1 - 10 with my stock answer of, "uh, maybe about a 7." So, what I'm attempting to do here is write down some loose specifications of the sort of instrument I'd like to see in the not-so-distant future, and perhaps comment a bit about reagents as we go along.

Lasers: I think the real key here, in terms of the number and wavelength of lasers, is options. If I had an unlimited budget, I think I'd probably put about 8 lasers on my cytometer (UV, Violet, Blue, Green, Yellow, Orange, Red, Far Red) pretty much covering the spectrum. I'd never dream of running all 8 lasers simultaneously, so they'd all need to have the ability to be shuttered on and off. I'm not a huge fan of turning lasers on and off constantly throughout the day, so I'd prefer to have them behind an electronic shutter. It's difficult to imagine purchasing an instrument with fewer than 4 lasers, but perhaps costs may force me to. I'd probably want to run as many as 5 lasers simultaneously, so we're aiming for 5 interrogation points. I'd also really like to have the ability to send lasers to different interrogation points. Most of the time, you'd probably not run UV excitable and Violet excitable dyes at the same time, so they could probably share a pinhole. But, in the instance where you would like to run them simultaneously, you'd probably want to split them to different pinholes and maybe even separate them by a pinhole or two. This would require some re-engineering of the way lasers are delivered to the flow cell, but I have a couple of ideas of how this might work, so it looks plausible. In terms of actual laser wavelength, that's to be determined. I'd need to weigh the merits of a 550nm laser versus a 561nm laser, etc... Regarding power, all I'd add is that I don't want to buy a 100mW laser to get 50mW at the point of interrogation.

Optics: Spectral overlap among fluorochromes excited off the same laser is to be avoided. So, it really doesn't make any sense to have more than 3 detectors off any one laser line. As you put more and more detectors on a single path, you have no choice but to break the light up into smaller and smaller bits, so by default you'll be compromising on photon collection; squeeze down the PE filter so you can run it, PETexasRed, and FITC all off the 488nm laser - this is absurd. You'd also want to stagger the emission filters so you're not looking at the same light from different paths that happen to excite off multiple lasers (think PerCP off the blue and PECy5.5 off the Green - change this to PECy5 off the green and PerCPCy5.5 off the blue). However, it DOES make sense to be able to detect lots of different fluorochromes off any single laser line. How can this be accomplished? Through quick change filters. For example, let's say we have 3 detectors off the Blue laser (SSC, FITC, PerCP, for example). I'd like to use that FITC detector for FITC, CFSE, GFP, mVenus, Aldefluor, Sytox Green, etc... Most people will have a 530/30 filter on their 'FITC' channel, but this may not be optimal for all the different 'green' fluors you may use. So, one option would be to use a wider band pass on that channel, say a 525/50. This is fine until I need to turn on my Green laser for excitation of some fluorescent proteins like mBanana. In this case I'd want to change my GFP filter to something like a 510/20, but then change it back when I'm not doing fluorescent protein work. Ideally, I'd like to tell the software which color combinations I'm using and have it adjust all the filters necessary to optimize fluorescence collection and minimize spectral overlap, but in the meantime, I want a system that has the ability to easily change filters, know which filter in in which detector, and have a place to store filters not currently being used so they don't get all scratched up and full of dust.

Electronics: I use to scoff at those who said they needed 5 and 6+ logs on their cytometer, but I'm coming around a little bit. I could easily see my next cytometer having at least 5 logs of dynamic range, but only if it has the right electronic components to fill those 5-6 decades. See this post for some ideas regarding that - Putting an End to the Log Wars. There's really no reason why our instruments should not have really fast processors that can do fine detail pulse processing. We're 11 years into the 21st century, yet we're using stuff developed in the 1980's. A great optical system is nothing without an equally great Electronics system.

Fluidics: In my eyes, Hydrodynamic focusing is still king (See edit below for clarification on how acoustic focusing is implemented specifically on the Attune ~~Acoustic focusing focuses the cells, but not the sample fluid so you end up picking up fluorescence from unbound fluors in the illumination volume~~ - this is the same issue with capillary systems), whether it's in a small chip or in a flow cell, however the sheath velocity going through the sensing area could be sped up to allow for higher event rates without increasing the size of the sample core stream. This, of course would require better electronics with much higher resolution to sample the short pulses and really good collection optics to collect as high a percent of emitted photons as possible, not just the small fraction that just happen to emit at 90 degrees to the incident light. I'd also caution against the desire to make super complicated fluidics systems that tend to break constantly (I'm looking at you FACSAria I and FACSCanto-A).

Software: Flow Cytometers are built by engineers, and it's usually the case that they find the engineer who knows most about writing software and say, "Let's get some software written to run this thing." There's usually not much usability testing, UI design thought, etc... The last batch of cytometers I've looked at have had a bit more polish on their software, so things look like they're headed in the right direction. The trend to borrow from MS Office and use ribbons all over the place is probably a safe bet. You'd have to assume Microsoft has done a bunch of usability testing, and if it's good enough for them, it's probably good enough for us. However, we're not word processing or making tables or even making presentations, we're adjusting hardware components using software tools, collecting data, and displaying that data on screen. So, in reality, we should be using a model that the everyday Jane Q. Researcher would be familiar with that performs a similar task. I'll throw out a couple of examples. I love the OSD (On Screen Display) on my Samsung LCD television. It allows me to easily get in, adjust settings like Color mode, Brightness, and Sound and get out all while not completely obstructing my view of the picture behind. Just change out Color mode, Brightness and Sound with Parameters, Voltage, and Compensation and switch picture with plots, and there you go. If you're a Photog, you probably use software like Aperture or Lightroom. These software tools allow for some pretty specific settings and adjustments but in a clean, easy-to-use interface. So, let's use these types of software to model our cytometer software after instead of a word processing software.

Reagents: I want lots of antibody choices, which is only going to be possible from a company that has ties to Research Institutions that make new antibodies and are willing to license them to companies for profit (eh-hem, our Monoclonal Antibody Facility has done and continues to do this on a regular basis). I also want them coupled to a wide range of fluors, especially the new ones like the brilliant violet. I want to be able to try before I commit, whether this be via a free sample, or a really inexpensive small aliquot. I'm not at all concerned about having reagents tied to my equipment, and I actually dislike that trend. I'm not going to buy a cytometer because some company made a canned "apoptosis kit" that works specifically for their instrument only to find out it's using Annexin V FITC and PI. The 5 questions I ask when finding an antibody. Do they have the antibody? Is it coupled to a range of fluors that work for my cytometer configuration? Does it fit in my budget? Are there multiple size options? Is there support information so I know it's going to work?

So, there you have it. How much am I willing to spend on this instrument? Well, I'm willing to buy as much instrument as I need. If I want a 2-laser, 6-color instrument, I think it has to be priced around $100K. If I want an 8-laser, 15-FL detector (5 pinholes x 3 detectors) with all the filters I need to look at 45 distinct fluorochromes, I'd say it'd have to be around $350K.

Edit: A comment above about acoustic focusing may be only partially correct. Although it is true that acoustic focusing is responsible for focusing the cells and not the sample core stream, this is not how it has been implemented in the Life Technologies Attune Cytometer. In fact, the Attune has both acoustic focusing for the cells and hydrodynamic focusing for the sample core stream. When utilizing the low flow rate on the Attune (25ul/min), you can achieve a significant amount of core stream tapering due to a narrowing of the entire stream from a cross-sectional area of 340um (where the cells are being focused by the acoustic wave form) to a 200um cuvette (where laser interrogation occurs). In this case, the constriction of the entire stream provides the hydrodynamic force needed to narrow the core stream. When running the sample at a higher flow rate, you'd increase the size of the core stream proportionally as is what happens on non-acoustically focused systems. However, in the case of the Attune, even at this higher volume flow rate, the cells still remain focused leading to better and more uniform illumination by the lasers.

Putting an end to the "Log Wars"

2011-06-15T13:49:00.000-05:00

A long time ago, in a laboratory far far away there was a lowly FACScan able to display data on a 4-log scale. Fast-forward to today, and you'll find some instruments with as many as 7 logs of scale. That's a huge improvement, right? Well, maybe not. The origin of the 4-log scale probably had more to do with the Analog-to-Digital Converters (ADC) being used than the technological needs of the science being done in the 80s. With the advancements in ADCs in other markets, flow cytometry manufacturers could now include converters with greater bit density and still provide a relatively affordable product. The standard for many years was the 10-bit ADC, which yields 1024 bins of resolution across the scale. Spreading these 1024 bins across a 4-log scale appears to give enough resolution while expanding to a reasonable range. After many years using these solid electronic components, BD completely redesigned the electronic system on its cell sorter (called the BD FACSVantage) to give us the FACSDiVa (or Digital Vantage) architecture. Now, instead of using traditional ADCs and log amplifiers, BD switched things up by using "high" speed Digital Signal Processors (DSPs) to directly digitize the analog pulse and then do log conversion using look-up tables. The DSPs converted the linear data at a bin density of 14-bit (16,384 bins) and when the data is log converted, it is upscaled to 18-bit (262144 bins). Now, with 18-bit data, they are able to display this data on a 5-log scale. The reason? Well, if I were forced to guess, I'd say it was a marketing decision to differentiate BD's new line of cytometers from it's old line as well as it's competitors. With this new 5-log data came with it the "picket fencing" phenomenon, which basically demonstrated that the 18-bit data (which was really 14-bit data) did not have enough bin resolution to display data properly in the 1st decade. The solution? Simple, hide the 1st decade and display decades 2 through 5 (right back at a 4-log scale). Because the BD instruments were so popular, other companies jumped on the bandwagon and thought, well if BD is doing 5-logs then we should do 6-logs or maybe 7-logs. And that's how we arrived here today, and now I'd like to show you why this is a bad thing.

Let me start with my conclusion first, and then show you how I arrived here. The figure to the right shows a minimun analog to digital conversion bit density for a given range of log scale. As you can see, if we wanted to display our data on a 5 log scale, we should have at least a 20-bit ADC. Side note - Bit(eff) means Effective Bit density, which basically takes into account that if you put a 20-bit ADC on your instrument, it probably doesn't actually perform at a full 20-bit. This is because there is some noise associated with the ADC, which limits the performance of the ADC. /Side note.

So, how did I arrive at this conclusion? Well first let me demonstrate that bit-density is important with an example. I created a mock data set of 3 Gaussian distributions (n=1000 data points for each) where the mean of the distributions and the SD were altered such that the populations were overlapping significantly. I then plotted these distributions on 4 histograms with different quantities of bin resolution ranging from 3-bit to 8-bit. It's important to remember that this is the exact same data set merely binned differently according to the available resolution. As you can see, the 3 populations are not at all discernable at the 3-bit range and it's not until we get to the 6-bit histogram that you can start to see the 3 different populations. Using this information, we can appreciate the importance of having sufficient bin density to resolve distributions from one another.

As an example to a system that might not have enough bin density, I display the following. Here we have a 20-bit ADC yielding over 1 million bins of resolution to spread across a 6-log scale. This may sound sufficient, but when we break it down per log, we see that in the first decade, where we have scale values of 1-10, we would only have 11 bins of resolution which would certainly lead to picket fencing and poor resolution of populations in that range. The Effective bins column shows an example where the noise of the ADC is such that our true bin resolution would be much less than the theoretical 20-bit.

Going through the process and crunching numbers for different scenarios, I conclude that ideally we would like to have on the order of 100s of bins of resolution in the 1st decade. So, in order to achieve that level on a 6-log scale, we'd actually need to have an 24-bit ADC. Now, the breakdown would be like what's shown below.

Take-home message: First of all, is a 6-log scale really necessary? For you the answer may be yes, but for most, probably not. The second question to ask your friendly sales representative is what sort of analog-to-digital conversion is done, and what the bit resolution of the converter is. It means nothing to have a 7-log scale displaying data from a 10-bit ADC. No matter how good the optics are you'll never be able to resolve dim populations from unstained cells. What really matters is having a really good optical system that has high speed, high density electronics that can display all the fine detail of your distributions. Find an instrument like that, and you have a winner.

Throw away your 8-peak beads...now.

2011-05-31T22:06:00.000-05:00

Why is it that each booth with a new piece of hardware I walked up to at CYTO had the same set of plots 'demonstrating' how 'sensitive' their instrument is? I can't tell you how disheartening it is after years of clamoring for a re-definition of instrument 'sensitivity' to see marketing materials littered with histograms proudly displaying 8 peaks. What does that actually mean? and Why do instrument manufacturers design instruments around this 'standard'?
Now, I can't totally claim innocence here. As you can see here, and here, I do use 8-peak beads as part of my panel of instrumentation tests. However, it's not the only test I run, and I use it more so to dismiss potential problems than single out an instrument that is performing particularly well. One of the best things 8-peaks can tell you about an instrument is the presence of background due to laser light bleed-through, or possibly a bad filter. 8-peaks are also pretty useful as an all-around alignment bead. Beyond that, there's not much you can infer from the resolution of 8-peak beads as to the ability of said instrument to resolve dimly stained cells labeled with a particular fluorochrome from unstained cells. For example, the ability to resolve 8-peaks in the FITC channel doesn't mean a whole lot as to its ability to resolve dim GFP signal from negative cells. Likewise, the inability to resolve 8-peaks doesn't necessarily mean that channel will perform poorly for a dim fluorescence signal. Let's look at an example.
I recently had the pleasure of evaluating the 8HT from EMD-Millipore (check out the full review here). For comparison's sake, I ran the same set of tubes on our Beckman Coulter Gallios. After collecting the data and doing the requisite comparisons, I noticed how closely the 8-peak data matched between the two, especially in the far red channel where we'd usually detect PECy7. The first slide below shows the 8-peak data for the PECy7 channel. The last 3 or 4 peaks are pretty much overlapping with one another. If we were using the 8-peaks as a metric of 'sensitivity' we'd probably conclude that these two instruments are pretty similar in their ability to resolve dimly fluorescent PECy7 stained cells from unstained cells, right?...right?

Not so fast. If we stain capture beads with a PECy7 antibody and look at the ability to resolve the 4 peaks that represent different levels of antigen density from each other as well as a blank bead, we can better assess (emphasis on the second syllable, please) the true 'sensitivity' of the two instruments. Looking below at this figure we can easily see that the Gallios is able to resolve PECy7 much better than the 8HT. This conclusion matches perfectly with real-world staining examples run on these instruments. It's obvious that if PECy7 was a pretty darn important conjugate for my panels, you know which instrument I'd be buying (if I could afford it, that is).

So, what's the take-home message here? Well, it's simple, 8-peak data cannot be used as a surrogate for how well an instrument will detect your panel of fluorochromes. What should you do? Again, simple: make sure YOU run YOUR favorite flavors of fluors on the instrument you're evaluating to give you an idea of how well it's set up for YOUR experiments. You can certainly do this by staining your cell samples of choice, or you could use a multi-peak capture bead to look at resolution. And if you want to quantitate this a bit more, you could extrapolate the peaks down to an area just above the blank bead to determine precisely how dim of a population you'd be able to resolve.

EMD-Millipore 8HT Review

2011-05-17T21:37:00.000-05:00

In my quest to find a mid-range cytometer to replace my ailing FACSCantos, I've come upon the 8HT from EMD-Millipore (whom I'll probably just call Millipore for now, or maybe even Guava at times). The 8HT is a 2 laser, 8-parameter cytometer (2 Scatter and 6 Fluorescence, setup in a 4-2 configuration). It represents the "top-of-the-line" instrument in the field of 8 cytometers that vary in their number of lasers, detectors and the absence/presence of a microtiter plate loader. The 8HT can accept either a 96 well plate or 10, 1.5mL microfuge tubes. There are also a set of 6 tube slots for various washing/rinsing purposes. There are a few unique features to this instrument which I'll briefly describe below before getting into the data.

All of the legacy Guava instruments and the new Millipore additions share the same microcapillary fluidics system. Whereas in flow-cell containing instruments, a sheath fluid is funneled through a conical shaped flow-cell where it creates the hydrodynamic force used to align the sample core stream through the laser interrogation point, here there is no sheath fluid. The microcapillary system is basically a clear straw through which the sample is drawn, and the microcapillary walls provide the physical barrier to align the sample, essentially replacing the sheath fluid. This basically means there's no PBS tanks to fill, and no giant waste container to dump down the sink. Also, since the sample core cannot expand in diameter to increase event rate like it would in a hydrodynamically focused system, the way in which you increase event rate is by literally increasing the speed at which the fluid is drawn through the capillary. With the capillary system, you do get a bonus in absolute counts for everything.

A second unique feature of the system is the modulated laser setup they've implemented to get around the need for separate pinholes and laser delays. Basically, the 488nm and 640nm laser lines are modulated out-of-phase with each other at a high frequency so that red fluorescence emitting from the cell while the blue laser is exciting is sensed separately from red fluorescence emitting from the cell while the red laser is exciting. I was pretty skeptical at first, but this works surprisingly well. For example, APC fluorescence was pretty well excluded from the Red1 channel (Blue excitation/Red emission). What makes this remarkable is that Red1 and Red2 are actually the same detector, but because of the modulation, the emission is able to be separated out cleanly.

The software, InCyte was pretty good. The thing that bugs me a little is the mere presence of the old Guava modules. The 8HT software, in general seems a bit schizophrenic. You can jump back and forth between a green background Guava software to a grey background InCyte. I think it would be less confusing if there were just one platform to use. The InCyte software is pretty capable on its own, so I can't see what's the use of the Guava software. I'm not one to use canned application-specific templates, so that makes anything with a green background pretty much useless. There are a few unique analysis tricks built-in as well, which show your data in a heat map-like graphic. However, probably my most favorite feature of the software is something fairly minor. To adjust the threshold on FSC, you can simply drag a red dotted line up and down the scale. There is no confusion on where the threshold is set.

So far, so good, right? Well, here's where things fall apart - Data. Like many of the other units I've tested in this range, their fluorescence resolution seems to be lacking. I'd put the 8HT pretty much on-par with the rest, but let's take a look at some figures. I ran my standard battery of tests including, Single Peak UltraRainbows (to get a glimpse at alignment via the CV), 8-peak rainbows (to assure a certain level of dynamic range and resolution), PI stained CENs (to look at linearity as well as well-to-well carryover), and my 'gold-standard' dim population resolution (using antibody stained capture beads). This time, to mix things up a bit, I chose to run the EXACT same samples on our Gallios, and just to make things absolutely fair, the samples were run on the 8HT first and then on the Gallios (just in case they started to deteriorate over time).

Single peak URFP, showing CVs of the fluorescence channels on the 8HT versus the Gallios.

8HT URFP Bead CV

Gallios URFP Bead CV

Next up, the ubiquitous 8-peak Rainbow Beads.

8HT 8-peak Rainbow Bead Resolution

Gallios 8-peak Rainbow Bead Resolution

PI Stained CENs: Here we notice a problem with the 8HT. My guess is that the unbound PI in solution is too much for the system to handle, and the detector is being swamped by light. Obviously, this could be alleviated by titrating the PI out, but as you can see below, the Gallios' baseline restoration has no problem with the unbound PI. If you think about it, in a capillary system, the amount of sample fluid that is illuminated along with the cell is much higher than in a hydrodynamically focused system running at a narrow core stream, so unbound fluorochrome in solution surrounding the cells has a huge impact on background. Keep this in mind when looking at the Dim population resolution data. Also, for carryover, I'm simply looking for cells in the subsequent well to have been stained by PI carried-over from the prior well. In this case, carryover appears to be less than 1:10,000.

For the Dim Population test, antibody binding beads stained with different fluorescently labelled CD4 antibodies were run on the 8HT at the 'very low' flow rate and the 'medium' flow rate. There are suppose to be 4 stained peaks (blue) and an unstained peak (grey). The lowest stained peak represents about 2500 bound antibodies (CD4 on Human PBMCs is about 50,000 binding sites). Here, we get a really great picture of the background issues. When looking at a blank bead by itself, the background is pretty low. However, once you have some fluorescence present, the background peaks merge together and offer no resolution. This effect is enhanced when you increase the flow rate. Again, the exact same samples are run on the Gallios.

So, as I've said many times, the convenience that these systems offer may be nice, but I'm not sure it outweighs the lack of fluorescence resolution. I'm sure things could be optimized so that this system would perform better, but I'll always come back to the fact that we don't run into these problems on our LSRII, Fortessa, Gallios, or even our Cantos, Caliburs and Scans. What's the difference? Hydrodyanmically focused streams with gel-coupled collection optics, high-quality/high-powered lasers, and bit-dense/fast sampling electronics. The quest continues...

Display Transformation and FlowJo: Confused? Read on.

2011-04-18T21:58:00.001-05:00

I'm not going to discuss the merits of transforming your log-scaled fluorescence flow cytometry data; I'll leave that to the professionals. What I will try to elucidate here is how I tweak the transformation using FlowJo (Mac version 9.3, sorry Windows people). For simplicity, I will use the factory default preferences to start. It's important to note that if you're on a shared computer, the preferences could have been changed, which may result in some funky things happening. It's probably best to get a set of preferences that you like, and save them so each time you can pull up your preferences. The preferences that matter for this are the Compensation section in the workspace tab (red box) and the "Define" button for 32-bit data, which will open the Options window for these type of data.

There are basically 3 things you can modify when adjusting the display transformation. They are the Number of decades, Additional negative display size and the Width basis. The Number of decades controls, to a large extent, how many decades of dynamic range is shown for events greater than zero. By default, this is 4.5--even if you export 5.5 decade data, use 4-4.5, otherwise too much visual space is devoted to the lowest decade. Additional negative display size controls, to a large extent, how much visual space to devote to events that have values less than zero. Since we are displaying data on a log scale, zero is not defined. So, there needs to be a way to display data around zero that makes sense. This is what's at the heart of the biexponential display. Near zero, the log scale becomes linear so that zero can be defined, and then goes back to log when safely in the negative realm. The number of channels around zero that are transformed into the linear realm is defined by the width basis. By default, FACSDiVa uses a width basis of -100, whereas FlowJo's default is -10.

So, how does this affect your data? Well let's take a look at an example. FITC and PE stained capture beads were run on a FACSCanto and exported in the FCS 3.0 format. No compensation was done at the instrument, and single stained controls were used to compensate the data in FlowJo. Below is the uncompensated data file as well as a compensated file using the defaults that were currently applied.

Looks pretty good, but let's take a look at some simple tweaks. For this, we'll go to the Platform menu, then Biexponential Transformation and then Manually Specify Transformation, which will bring open the window to edit the transformation settings.

Again, we have the option to change the width basis, Positive decades and Additional Negative Decades. Let's assume we're not going to change the Positive decades, so we'll focus on what the width basis and negative decades will make. Below are the plots shown at each of the width basis presets. As you can see, the main affect is squishing the data closer together around the zero point. Good to get data off the axis, but you can easily take it too far and end up reducing your ability to resolve dimly stained cells from unstained cells.

The question then is how much transformation do you apply? One strategy that I like to employ is to try and visualize this better with a contour. The goal is to remove the bimodal-like profile of the populations as they cross the zero point. Once I'm able to do that, I then increase the amount of negative log space so that most of the data is not on the axis. For example, below I show a -10 width basis, 1 additional negative log in a contour plot. In the brightest peaks, it is easy to see a pronounced dumbbell shaped population straddling the zero point. If I modify the width basis a bit to get rid of the dumbbell shape, and then reduce the additional negative space to remove extraneous white space, I get a profile like the one on the right. Notice that each axis is done separately and can have different width basis and negative logs to achieve the best transformation.

Once all is said and done, I now have a well transformed plot that is worthy of publication. Below is the original uncompensated plot, the default transformation, and the modified transformation.

A few tweaks and a bit of trial and error is all you need to get visually pleasing plots that will actually help you make better decisions in terms of region drawing and data interpretation. So, please feel free to play around with these settings and see how well you can transform.

Sort Schedule Changes: The Who, What and Why.

2011-03-31T12:11:00.000-05:00

Let's start at the beginning. Back in the days of the FACStar Plus and water cooled gas lasers, we would begin accepting sort reservations at 10AM. This allowed us to get in at 9AM, warm up the lasers for 30 minutes, get the fluidics going, and have everything running smoothly by 10AM. We typically did 1 or (on a good day) 2 sorts, which lasted 2-4 hours going at a rate of about 5000 events per second. Adding the MoFlo in 2001 allowed us greater throughput by allowing us to run at 10's of thousands of cells per second, but we still needed a good amount of time to get started in the morning so the I-90s could warm up and the fluidics could stabilize.

As word got out about sorting applications in general, the sorters became busier and busier; so much so, that we traded in the old FACStar Plus, and traded up to the FACSAria. Around the same time, we retired our water cooled gas lasers, and outfitted the MoFlo with the solid state Lyt-200 from iCyt. The ease of setup on the Aria, and the nearly instant-on of the solid state lasers made getting started in the morning much easier. Slowly but surely, we allowed earlier and earlier sorts until we officially added the 9AM - 10AM slot on both sorters. The increased capacity kept up with demand for a period of time, but soon even running both sorters 9AM - 5PM wasn't enough. We also saw that the afternoon slots were filled weeks in advance, and the unfortunate few who had to book sorts last minute were stuck with the 9AM slot, usually forcing them to get started in the wee hours of the morning. In an attempt to solve this problem, and position ourselves for an S-10 application for a 3rd sorter, we added a 5PM - 8PM "late night" sorting slot on Tuesdays and Thursdays, and had one of the operators work a 12PM - 8PM shift on those days. These slots were gobbled up instantly, and soon became a favored slot amongst the users. Still struggling with overall capacity, the cancer center stepped up and helped us purchase another sorter, the FACSAria II. So, now we had 3 sorters available from 9AM - 5PM Monday thru Friday, and the FACSAria II available an additional 3 hours on Tuesday and Thursday evening. This setup served us very well for a couple of years. Along the way, we still saw that afternoons filled up very quickly, and the 9AM slot was still being used for last minute sorts, or GFP cell line sorts.

So, we decided to take a look at usage more closely, and query our user group about the possibility of opening more 5PM - 8PM slots. Of course everyone thought the idea of more evening slots was great, but we needed some data to support this change. The figure below shows the frequency in a year of a sorter being used at each hour of the day. It is separated out for each sorter. The blue line shows what percentage of available time the sorters (as a whole) are being used. As expected, the afternoon slots are used 70% of the time, while the 9AM slot is reserved only 25% of the time. Also, the 5PM - 7PM slots are used 60% of the time, even though the actual number of sorts taking place appears low. The 25% number at 9AM is also a bit misleading. This represents that hour being booked, but I can tell you, it is very rare for someone who books a sort from 9AM until 11AM to actually show up at 9AM, and the longer that reservation is, the later they show up. So, the actual usage of the 9AM hour is probably much lower. I'd estimate it at 10% or so. So, instead of sitting around waiting for the 9AM sort to show up, we could utilize that time better by getting QC done on our army of bench-top analyzers, or allow for more stabilization time of the sorters to get even better results. So, beginning in April, we will have new sorting hours as follows: M - F 10AM -5PM, and additionally on one sorter, 5PM - 8PM M - Th (no one wants to sort until 8PM on Friday night!). The number of available hours per week is still about the same. Previously, we offered 121 hours of capacity each week, and now we're offering 117 hours. Our actual weekly usage is about 70% of that anyhow, so we feel the new hours are plenty for the current demand. In addition to all this, we have been aggressively pursuing users who are frequent FACSAria users to get trained and run their own sorts. We have half a dozen users who already do this on a regular basis and will be tapping a few more competent users in the near future. We feel this, along with the increased evening hours, and the removal of the 9AM slot, will maximize the efficiency of our sorters and allow more convenient access for our users.

Getting in tune with the Attune

2011-03-23T22:33:00.000-05:00

EDIT 2: I've met with some folks from Life Technologies and they feel the instrument that was set up in my lab for demo was not properly aligned, and so I've agreed to give it another try. I'm not sure if I'll use the Violet/Blue combo or the Red/Blue system, but either way, I'll post the results and reference back to this post when I have them.

EDIT: Regarding the statement below in the Optics section, "I have not received information that I requested yet, but the 'PMTs' do not appear to be 'PMTs'." I have heard back and there are, in fact, PMTs in the system. It's just not obviously visible when you simply open the hood. They are the Hamamatsu H10720 Series, and in the far red channel, they are using the Red sensitive "-20's"

You've probably heard about the novel approach to sample focusing on the Attune, and you've probably even read through some of the marketing materials distributed by Applied Biosystems (Life Technologies). So I won't go through all those features in detail, but I will share my impressions and tell you what it means for your data in the end.

Briefly, the Attune is a 2-laser, 6-fluorescence detector small-footprint cytometer. The fluidics are syringe pump driven (as opposed to pressure-driven), and it's in the focusing of the cells through the laser interrogation point that is unique. Instead of hydrodynamic focusing applied by a sheath fluid (such as PBS), the Attune uses an acoustic wave form to push the cells into the center of the stream, forcing them to line up in single file. One advantage of this is the small amount of sheath buffer you need to put into the system (roughly 1L per day compared to 6-8L for a hydrodynamically focused system). Another benefit is the ability to push through large quantities of sample fluid per unit time. Whereas a pressurized, hydrodynamically focused system might top out at about 200ul/minute, the Attune can achieve volume flow rates of 1000ul/minute. It can also maintain the CV of the fluorescence signals even at high flow rates. Lastly, the system can actually change how fast the events are passing through the laser interrogation point. They have a standard, and high-sensitivity setting, which tells the system to pass them through at the normal rate, or at a slower rate, respectively. The idea is that if the cells pass through the lasers more slowly, the fluorochromes will emit more photons, allowing you to detect lower amounts of fluorescence.

Optically, the system has a 20mW 488nm laser and a 50mW 405nm laser. There are no other options for laser lines at this point. 3 of the fluorescence detectors are dedicated to the blue laser and 3 to the violet laser, and they're set up for FITC, PE, PerCP/PECy7, and PacBlue, Qdot 525, and Qdot 605/PacOrange. The filters are easily changeable, and there's even a storage section for spare filters (how nice!).

I'll try to break up my comments by instrument components. I will say, many of the complaints I had regarding the software were said to be fixed in an upcoming release, but I will give my opinions based on the instrument I evaluated (03/09/11). Also note that the instrument was installed by a field service engineer and passed their performance and tracking protocol (similar to CS&T a la BD).

Fluidics:
1. Being a syringe pump system, you need to specify how much volume you wish to analyze before you even put on your sample. Of course, you can always add to your sample acquisition, but it doesn't have the 'free-run' capabilities that a pressurized system has.
2. If you tell it to take up 500ul of sample, and then you get enough data, or you want to just take your tube off, the remaining sample in the line goes to waste, and not back to your tube. So you could foresee a possible loss of sample in some situations.
3. The dead volume is huge! 200ul of your sample is taken regardless of how much sample you wish to analyze. So, if you want to analyze 100ul, you need to have at least 300ul of volume in your sample.
4. It was really nice being able to put on different sizes of tubes and not have to worry if they're the right kind or cracked.
5. As advertised, the CV's are really low even going at high sample throughput rates.
6. Although they claim they can run at rates of up to 20,000 events per second, there is no report of the coincident rate going at that speed. A high abort rate would obviously have an impact on detecting and counting rare events. Even on our systems with zero "dead time" we still see coincident events to some degree.
7. Although in theory we'd expect the high sensitivity setting (i.e. slowing the cells down in the interrogation point) to increase resolution, my tests did not show any improvement with stained beads (data not shown). It's likely that the components that contribute to autofluorescence also increase their intensity so the separation between positive and negative doesn't really change.
8. When you tell the system to collect 2mL of sample volume, it does so in (for example) 500ul 'draws' from the tube, i.e. 4 draws from the sample tube. The reason is that they don't want to have all that volume sitting in the tubing waiting to go to the flow cell. The result is that you end up having these pauses in acquisition while the system refills the tubing. Because of this, when you start collecting your sample again, the core stream has not completely stabilized leading to a decrease in fluorescence and a widening of the CV (Data Below). In order to get rid of this data, which will surely mess up your analysis, you'd have to go to each point where it drew more sample and exclude the first few seconds of collection until the stream has stabilized. It's an easy enough fix to make the software ignore data being collected for a few seconds every time it draws, and hopefully they'll implement that fix.

Software:
1. IT'S FREE!!!!!!!!
2. The overall look of the software steals a page from Microsoft Office, with a prominent ribbon at the top for common tasks, and lots of right-click functionality. Users should feel comfortable here.
3. Similar to FACSDiVa software, there is a Experiment Explorer (Browser) that displays your experiment, all the tubes in the experiment, and the settings/plots saved within. There was some confusion in terms of things being applied globally (to the entire experiment) or to an individual tube. Again, similar to DiVa, it's not always obvious if something is applied globally/individually, and how to change that status easily. Lastly, it also resembled the Gallios software in that if you made a change to the plots/regions, you had to click save or else you'd lose those changes when you switched tubes.
4. Gating was somewhat cumbersome. Changing hierarchy of the gating structure after you've created gates was difficult and not intuitive.
5. Displaying large numbers of events was slow and jerky. There was this weird need to press 'F5' to refresh plots with lots of data point.

Optics:
1. I have not received information that I requested yet, but the 'PMTs' do not appear to be 'PMTs'. There is definitely not a vacuum tube with anode, dynodes and cathode like you'd expect to see. It looks like some sort of silicon diode, but like I said, I haven't heard yet. This could contribute somewhat to the performance I observed.
2. I really appreciate the extra slots for filters on the instrument. I'm a big believer in spare filters, but I can never seem to keep track of them (just look in our MoFlo room).
3. Although the laser powers seem appropriate, there was no mention of how much of that light actually makes it to the flow cell.

Electronics:
1. Testing of linearity across the 6-log scale was good (CEN data below)
2. 23-bit ADC conversion yield 8.4million channels across 6 decades, which generates approximately 76 bins in the 1st decade. This seems pretty good, and would allow for resolution of closely related populations on the low end of the scale, but that's not really what I saw. I was able to see some 'picket fencing' which tells me that the noise of the ADC makes the bit conversion probably closer to 20 bit, which yield about 9 channels of resolution in the first decade. This is speculation on my part since I do not know (yet) what ADC they're using, and what the noise level of that ADC is.

Data: For this I ran my standard battery of tests, the results of which can be found below.

8-peaks: Testing the overall alignment and diagnose background problems due to laser light.
Assessment: 8 peak resolution off the blue laser was not as good as other 'full-size' cytometers (i.e. FACSCanto-II, or Gallios)

CEN Linearity: Using PI stained (and DAPI stained, but not shown) CENs.

Assessment: Linearity was good along all of the scale tested.

Dim Population Resolution: Here we have Antibody binding beads from Bangs stained with a FITC, PE or PacBlue antibody to demonstrate the ability to resolve increasingly dimmer fluorescence from background. The more separated the peaks, and the further from the blank the better the resolution.

Assessment: The FITC channel (BL1) seemed to be the worst of the 3 tested. The PE channel was ok, and the PacBlue Channel (VL1) was pretty good. To see comparable data on another instrument, check out the same data on the FACSCanto-II

Stream Stability: For this, I tried to have it draw sample multiple times while looking at PI fluorescence in linear.

Assessment: As you can see, the fluorescence dips down every time this fluidics draws more sample. This leads to an overall higher CV if you don't time gate the data. You can easily see this by comparing the purple box at the beginning of acquisition post draw and the green box that's mid-way through that collection.

Final Thoughts: Like many of the smaller sized cytometers, the Attune will probably handle most of the general types of applications, but will not compare to high-end cytometers in its ability to detect dimly stained populations, even using its High Sensitivity mode. The real advantage is its ability to push large amounts of volume of a relatively dilute sample through the instrument without increasing CVs of the populations. For example, a lyse/no-wash rare event blood tube. In most every other case, we would simply concentrate the sample by spinning it down, and run it in a similar amount of time as what can be achieved on the Attune. The overall look and feel of the software is fairly polished. If they're able to correct a few major things (like return unused sample, not store data during stream restabilization, allow for a quicker refresh of data on the screen, and get rid of the F5 necessity) with some software changes, it will be a much better instrument. The lack of a red laser, or laser options is a real hurdle for many people, especially those with a large stock of APC and APCCy7 antibodies, and could be a deal-breaker for many labs. It would also be nice to have a higher powered 488nm option as well. This, ~~along with the whole 'PMT' issue~~ might lead to better resolution in the FITC channel, which was especially poor.

Just when I thought I was out...they pull me back in.

2011-03-10T16:07:00.000-06:00

I'm speaking, of course, about BD and the FACSCanto platform. Now, if you know me and have talked to me about flow cytometers, you know I haven't been too kind to BD and the FACSCanto-A. We have, and continue to, battled with problems on these instruments. It doesn't help much that we have hundreds of users running all sorts of who-knows-what through the instruments. But then again, our LSRII's never break-down, our 15 year-old FACScan and 10 year-old FACSCalibur never break on us and they get just as much use. Recently, however, I happen to have the privilege of running a FACSCanto II with an HTS. We've had it in the lab for a few weeks, so I've been playing around with it trying my darndest to break it, without success. It's definitely not a fair comparison as far as the use and abuse our other cytometers deal with, but I've tried to run some unfiltered chunky samples on it a few times, and it recovers well. The HTS has been running as well as could be expected; so well, that we're now putting one on our Fortessa. There are two other things on the Canto-II that have really caught my attention. The first thing is the overall fluorescence sensitivity and resolution. It certainly ranks among the top instruments I've ever tested. I ran my standard battery of tests on the instrument (dim population resolution, linearity, dynamic range, and precision) and the FACSCanto-II excelled in all respects. I'll highlight a few of these tests with some figures below. Before that, I want to document just how awesome it is to be able to put on a regular 12x75 tube with about 100ul, and be able to analyze almost all 100ul out of the tube. The way that the lever pushes the tube all the way up so that the probe is nearly hitting the bottom is awesome. That, combined with the absence of a DCM sleeve on the SIT makes this possible. Dead volume is about as minimal as can be for a tube loader.

Figure 1 below simply shows some antibody binding beads from Bangs stained with a CD4 antibody coupled to either FITC, PE, or eFluor450. The lowest stained peak represents a binding capacity of ~3000 antigen. To put that into perspective, CD4 on human Tcells is expressed at about 50,000 antigen. So, you can see that something with 10-fold less antigen density can easily be separated from background.

Figure 2 is simply displaying 8-peak bead data.

And Figure 3 is showing PI stained CEN's to show precision (% CV of the 1st peak) and linearity peak-to-peak ratios.

A 19.2V Drill with every sorter purchase?

2011-02-07T09:35:00.001-06:00

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.

The Beckman Coulter Gallios and Translational Research at UCFlow

2011-01-07T11:38:00.000-06:00

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.

Compensation is infiltrating my dreams.

2010-11-09T09:24:00.000-06:00

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.

MoFlo Upgraded to XDP, plus a couple new laser lines.

2010-10-01T11:38:00.000-05:00

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 -

The remains of the MoFlo after the tear-down

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.

The upgraded MoFlo XDP in all its polished glory!

Follow us on Facebook!!

2010-09-23T11:49:00.000-05:00

The flow lab created its very own group on Facebook where you'll find the latest news about us, information on our instruments and discussions on the up-to-date events in the fantastic world of flow.

Find the University of Chicago Flow Cytometry Core Facility group and feel free to leave any comments and questions you might have.

GLIIFCA Core Manager Meeting Preview - September 24, 2010

2010-09-22T22:19:00.002-05:00

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.

Flow cytometry leads to everything!

2010-09-16T11:11:00.000-05:00

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.