Thursday, March 26, 2015

Index Sorting - From FACSDiVa to FlowJo

We recently upgraded our FACSAria to FACSDiVa 8 running on Windows 7 primarily for the ability to do index sorting. Getting used to a brand new set of DiVa issues and quirks has been difficult, but we soldiered on nonetheless. After scouring the web for resources on both index sorting and analyzing index sorting data outside of FACSDiVa, I decided to compile all the resources in one place. They are out there, it's just a pain to jump around to various sites trying to compile all the information together. I've done the leg work already, so read on to get the info. Of course, I'm sure there are more elegant ways of doing this in other programs or even in FlowJo, but I needed this info yesterday, so I'm documenting it here for future reference. 

Figuring out index sorting in FACSDiVa. You may think index sorting is no more than checking a box in FACSDiVa, but there are enough one-off situations that arise that it really warrants a separate FAQ. There are two resources that are quite helpful in figuring this part out. The first, oddly enough, is BD's very own Index Sorting Manual (<-- fixed bad link), which comes as an addendum to the FACSDiVa software manual and may not even be installed on your computer or available for download from BD's website. I only came upon this after our BD service engineer sent me a copy of it. The second resource is a document presented at GLIIFCA 2014 by Matt Cochran (University of Rochester), in which he outlines some of his tips and tricks for working with index sorting in FACSDiVa 8. 

So, let's assume you figure out how to successfully perform an index sort in FACSDiVa. You should have a Pre-sort FCS file of your entire population, and an Index sort "tube" for each plate you ran. You can export both (or all) of these as FCS files. There is a decent interface for looking at your index sort plate information within FACSDiVa, but if you're use to doing all your analysis is FlowJo, you probably want to bring that data over at some point. And here's the fun part.

Analyzing index sorting data in FlowJo. I have an application where a user is index sorting based on a range of FITC intensities. The resulting plate will be a mix of FITC low and FITC high clones. The goal of index sorting, in this case, is to retain the original FITC intensity information for each well after the sort. What follows below is A method (not THE method) I stumbled upon to go from an index sort file from FACSDiVa to Figure 1 below. I'd really love for someone to tell me there is a way easier way to do this in FlowJo.
Figure 1. Heatmap analysis of index sorting file.

Figure 2. Running the initial script to create 96 populations
Step 1: Use the Script Editor index sorting example from the Daily Dongle Blog (or see Addendum below regarding the method in Version 9). You simply copy and paste the script starting with  /** --- Iterate samples --- **/ all the way through gate.update(); } and paste it into the script editor window (under the tools tab of the ribbon) in version 10.0.7 (if you have access to the 10.0.8beta version, I would do this step in that'll see why later).  Highlight your index sorting file in the workspace and click the run button in the script editor window. You should now have 96 populations under your index sorting file. If you end up with a bunch of "-" where it usually says the number of cells in each population, click the refresh button at the top of the workspace window and then it'll show you that there is 1 cell per region (Figure 2). 

Step 2: The next step is to export each of these populations as its own FCS file. In essence creating 96 FCS files. The problem here is that you can do the initial index sorting script in version 10.0.7, but you can't do the export to 96 FCS files in 10.0.7 for the Mac (I think you can do this in the windows version, but I'm not sure). You can do the export to 96 separate files in Mac version 9.8.3, but you can't do the initial script in 9.8.3. So, if you can do this all in 10.0.8beta, that's your best bet (or on windows). So, in 10.0.8b, you can highlight all the populations and choose export (right click or within the File tab in the ribbon) and export this as 96 FCS files. 

Step 3: Using the plate layout to create a heatmap. The last step is to load the 96 FCS files into FlowJo v10.x.x and assign the Well ID keyword to each of the files corresponding to their position on the plate. Now, the files are in chronological order going across and then down (in serpentine fashion). So all you have to do is add the Well ID keyword as a column and copy and paste a list of Well IDs (A1 - H12 in serpentine fashion) from a spreadsheet. BUT WAIT, THERE's MORE! If you're doing this on a Mac, this post from the Daily Dongle states that since Mac Excel copies data in the ANSI format you won't be able to paste into FlowJo, which only reads the Unicode format. To get around this, create the Well ID list in Google Sheets and copy and paste from there (Google Sheets copies data in Unicode format). Now that you have a Well ID associated with each of the files you can use this link to in FlowJo's documentation to set up a heatmap of your index sorting data.

And there you have it. Please leave a comment below with your preferred method of analyzing index sorting data using whatever software you like. 

Addendum #1: Using the script is somewhat cumbersome. Thanks to Helene Dujardin (from HCD Bioexperts) for the tip below:

"There is another way in version 9, has there is an option for index sort analysis.  Select your sample and go to the menu Platform/ Event number gate / Create Indexed sort gates. It will directly create a gate for each of your well. Each gate name will be the corresponding well ID.

You can then export each of your gate as a new fcs file also with version 9. Your exported fcs file name will include the well ID if your original fcs file name is not too long (you can change it by changing the $FIL keyword)."

Addendum #2: Using the methods outlined in Addendum #1, I'll add one more point of interest. When you export the Index FCS file from FACSDiVa, you might get a really long name (Specimen_001_Index_Tube_001.fcs). FlowJo v9 freezes when you try and export all the regions as FCS files, so you'll need to rename the files after you import the parent into FlowJo. I've been renaming them (CMD+R shortcut) INDX_1, INDX2, etc... Now, when you export the regions as FCS files they'll be labeled INDX_1_A01, etc...

Wednesday, December 31, 2014

Flow Cytometry Core Facility New Year's Resolutions

It's that time of year again when the gyms are packed and weight-loss commercials air continuously.This year, why not turn you attention towards your core facility and come up with some resolutions the whole lab can take part in. The best part is you'll have help from the rest of your lab mates to keep you on task.

So, just as we do with our personal lives, allow me to present an ambitious 10 resolutions for the UCFlow core facility. Presented in no particular order, I give you:

  1. I'd love to devote more time towards taking better care of our instruments, in terms of routine maintenance and a more streamlined QA process across the board.
  2. Do a better job getting administrative tasks like billing/invoicing/usage tracking/usage analysis done on-time and with greater regularity.
  3. It's always nice to see how the work done in the core fits into the bigger picture, so I would like to go to more of my user's talks on campus.
  4. It's pretty clear data analysis is a hot topic these days, so I want to focus more attention on complex data analysis solutions for users (is R worth it?, try more advanced stuff in Cytobank or FlowJo?, etc...)
  5. Who can't use more/new instruments. You'll get none of the instruments you don't write a grant for. I think I need to be more aggressive in my pursuit of new funding sources for instrumentation.
  6. Blog more often (a perennial resolution for me).
  7. I'm convinced that the Hangouts on Air that we do in the Cytometry Community on Google+ are super useful, and so I'd like to turn that into a more regular thing. 
  8. I've always thought that eventually core facilities would collapse into each other to create mega technology centers. But, before that happens, I would like to start by increasing interactions with other core facilities on campus to see what they're doing and what's new in technology in other fields.
  9. It use to be the rule in our core that if you went to a meeting, you had to present something. I haven't been as faithful to that rule as I would like, so I'm bringing it back.
  10. Of course this last one happens all the time, but I would like to focus some attention on re-evaluating facility costs with greater scrutiny to determine where reductions can be made.
Well, there you have it. I just hope I'll be able to hold onto these longer than my annual attempts to get back "in shape." How about you? Any resolutions you'd like to add for your core facility? Leave a comment.

Friday, December 19, 2014

Core Facility Acknowledgment Accounting 101 - How to make sure your work is being recognized.
BioTechniques Article on SRL Attribution
A recent article in Biotechniques has spurred some interesting discussions in the Academic Core Facility (or as we cytometry cores like to call them, Shared Resource Laboratories - SRLs) world. The gist of the article states that all too often core facilities are not properly acknowledged in publications that clearly are using the services provided by their institutional cores. The flip-side of this argument is that investigators are already paying for the services rendered so that fee is essentially all the "acknowledgment" that is required. However, since many times core facilities are partially funded by government agencies, the services (and more accurately the service recharge rates) are being subsidized. Therefore, the payment isn't payment enough.

Whether you agree or disagree with this basic tenet is really beyond the scope of this post. What I'd like to share here is my way of fostering the proper relationship with my users such that they feel compelled to acknowledge the excellent work of the core instead of feeling obligated to do so.

What follows is basically a three-part approach to accomplishing the goal of being acknowledged as a core facility in publications that utilize your services. The reason you may wish to do this could vary, but likely involve justification of your core facility's existence to your institution's administrators or various "Centers" you may receive funding from. For example, as part of the University of Chicago's designation as a Comprehensive Cancer Center from the NCI, we must keep track of cancer-related publications that utilize our core facility. So, obviously it would be easiest for us to search PubMed for the inclusion of our core facility's name or even the cancer center support grant number in the reference. However, many times our facility is omitted from the acknowledgement section of the publication. To help modify this behavior, we need to first find the publications, then organize them, and lastly reach out to our authors/users to help them understand why acknowledgements are important. Here are these steps.

Part 1 - Finding publications that should designate attributed to your core facility's work.

Fig. 1 - Keywords to find references based on your core's services.
You can use the "Saved Searches" functionality within PubMed to find relevant articles and have them emailed to you directly as soon as something meets the search criteria. There's already a good tutorial on PubMed that will walk you through the steps, so I won't go into that in great detail, but let me summarize my steps.

Figure 2. Part #1 of search yields over 168,000 results.
I jump right into the Advanced Search Builder in PubMed using various keywords for different parts of the search structure. For example, I limit the search results to an affiliation of University of Chicago. There are a few external users that I'd like to track as well, but I put them in a separate search. The first part is to put in keywords based on any part of the text that your users may use to describe what they did in your facility. Remember, many users refer to any part of flow cytometry as "FACS" so you'll want to make that part of your search criteria. Figure 1 shows you some of the ones I use (note the use of 'Or' boolean to search on any of these keywords).
Figure 3. Search restricted to affiliation of University of Chicago

Next I use the "Add to history" link near the search to hold onto those search results temporarily (Figure 2).

I click the "Add" link next to search #1 to add these 168,000+ results back into the builder, and then refine the search by using the "And" boolean and restricting the "Affiliation" field with 'University of Chicago (Figure 3.)

Figure 4. Search based on keywords, affiliation, date range
You can further refine the search based on Date ranges or excluding reviews or a bunch of other search criteria using the same strategy (Add to history, then add those results back to the Builder and refine again). I find this method of going back and forth between the history table and the builder easier than trying to put everything into one complex boolean structure. Click search to view your results (Figure 4).

Once you've created your search criteria and confirmed that it is giving you what you've intended, you'll want to save the search, using the "Save search" link below the search box. Figure 5 shows you some of the options available for setting up the saved search. Note that you'll need a PubMed profile to set this up, so the first time you try and save a search, it'll ask you to create an account. Here, I've chosen to send me an email
Figure 5. Saving the search and setting up email digest.
weekly on Mondays (when I'm likely to have free time) so I can review the new references. I've also placed some text (or even a custom #) so that I can filter my email properly and it doesn't get lost amongst the email clutter. I save the search and wait for the emails. By the way, you can now set up all sorts of notification. For example, I've recently been doing a lot of microparticle stuff, so I have a separate digest setup to send me email notifications of new publications using flow/image cytometry to analyze microparticles (or microvesicles or micro particles, etc...)

Part 2 - Organize references and tag them to easily create reports later.

In part 2, my goal is to receive these email notifications, skim through the publication and then find a way to organize the references neatly and efficiently.

Figure 6. Email notification from My NCBI
The emails arrive in my inbox on Monday mornings as references become available. If there are no new references, you will not get an email. Figure 6 shows an example of what this email looks like.

Next, I follow the link, and read through the manuscript to ensure the work being reported was in fact from my core. If I'm unsure, I can always ask the author, but I tend to recognize work done on my instruments.

Figure 7. One-click add to Zotero button in URL bar
One thing that becomes evident is you need to have a way to manage all these references. There are a ton of ways to do this from the most rudimentary word doc or spreadsheet to sophisticated software management tools. The tool I like for this part is Zotero. It's similar in function to things like Endnote or Mendeley, but it's basically an organization tool for references. The part I like most about Zotero is that there's a Chrome extension that allows one-click adding of references to my database (Figure 7). Plus it will go out and find the PDF of the full-text reference and store that locally as well (when available). It lives in the cloud and can be accessed anywhere. 

Figure 8. Zotero Organizing tool for references (running on Mac)
Once in Zotero (Figure 8), I can add tags to the references to help organize them further. I like to tag things by services used (Cell Sorting vs. Analyzer Usage vs. Other things), Instrument referenced (e.g. FACSAria), Whether this could be used for my Cancer Center grant renewal (UCCCC), and other informative tags. Then, down the road when I need to pull up some justification for a new sorter, I can include a list of publications that utilized the cell sorting service or maybe even a specific sorter.

This makes organizing and searching through references a breeze.

Figure 9. Thanks for the acknowledgement
Part 3 - Compel investigators to acknowledge your core facility.

Now comes the hard part. How to suggest to your facility users that they should be acknowledging your core without sounding like a jerk.

As I'm skimming references, I'll quickly jump to the acknowledgement section and check for recognition of the core, or perhaps individual members of the core (either is fine with me). If the user does acknowledge the core, I make sure to send them an email thanking them for doing so. This positive reinforcement goes a long way toward ensuring this type of action recurs in the future. I also explain why it's important to us that the core be acknowledged. An example email is shown in Figure 9. Of course congratulating them on a job well done can only help to sweeten the deal.

Figure 10. Maybe next time...?
If I see there is no mention of the core in the acknowledgements or methods section, I'll send a similarly positive email, but ask them to consider acknowledging us in the future. I make sure to include some example text of what I would like them to say, as well as send them a link to the example text on our web site (Figure 10).

Of course, you can save these emails as templates and simply change the name and journal to personalize them.

The responses I've received from these emails has been tremendous. I think they are both appreciative of the recognition of their work as well as understanding of the needs of the core to be recognized.

We all understand the need for metrics such as publications and their importance in validating the success of core facilities. However, instead of taking a passive approach and hoping people read your web site asking to be acknowledged, the method proposed here takes a proactive approach that has already increased the desired result.

PubMed is pretty comprehensive, but there could be other sources for finding work being discussed that should point back to your core facility. Magazine articles, intra-institutional articles or highlights, blog posts, etc... all should be explored and stored. You can use a series of other rss feeds or Google search alerts to help you find this information too. Asking a PI to mention the core facility in an intra-institutional newsletter is certainly within your purview.

Happy Hunting!

Thursday, September 4, 2014

A First Look at the Beckman Coulter CytoFLEX - Strong Performance in a Small Box

Over the past few years, we've been inundated with small, inexpensive cytometers with the promise that they can perform as well as the big boys. Up until now, I would have told you not to waste your time... up until now.

In 2013, an unknown company called Xitogen set up a booth at the annual CYTO conference. Before long, there was a buzz racing through the exhibit floor aisles of a flow cytometer starting at ~US$25,000 (1 laser, 2 colors). The Chinese company, headquartered in the Suzhou Industrial Park, set out to provide an alternative for Chinese researchers to acquire affordable flow cytometry instrumentation without having to deal with overpriced imported hardware from the big players. With U.S. zero install base, and zero user-generated data, CYTO 2013 came and went, and the buzz surrounding Xitogen died out. It was pretty obvious the better known cytometer manufacturers would be taking a look at the company for a possible acquisition, and in April of 2014, Beckman Coulter announced they would purchase Xitogen for an undisclosed amount of money.  The acquisition was finalized in June 2014. At CYTO2014, Beckman Coulter revealed the re-branded instrument now called CytoFLEX.

I had the chance to spend about a month with the CytoFLEX and what follows are some of my thoughts about the key features, successes and failures of this instrument.

General Technical Specs:

The CytoFLEX came to me as a 3 laser system including a 50mW 488nm laser, a 55mW 640nm
Beckman Coulter CytoFLEX Analyzer
laser, and a 93mW 405nm laser. The system also has 9 fluorescence channels in a 4-3-2 configuration, respectively. In addition, there are 3 light scatter parameters, the typical blue laser scatter yielding forward and side scatter, and an additional side scatter parameter off the 405nm laser. Pulse height and area are collected for all parameters, and a width signal can be selected for any one of the parameters. The fluidics system is controlled through peristaltic pumps for both the sheath and sample lines, and the sample volume flow rates can range from 10ul/min up to 240ul/min with 10, 30, and 60 ul/min presets (referred to as Low, Med, and Hi, respectively). A single tube holder with built-in backflush loads samples into the instrument one-at-a-time, and the hardware is controlled by the bare bones, but highly functional CytExpert acquisition software.

The system that is suppose to ship some time in October will be configurable with 3 spatially separated lasers (with a 4th coming soon?), with a variety of laser options and colors available.  The base configuration should include 3 lasers, and 13-colors in a 5-5-3 config (violet, blue, red, respectively).

Look and feel:

A look inside the CytoFLEX revealing lots of unused space.
The instrument itself is quite small, fitting roughly into a 40cm cube, but even the box itself seems to be too big for whats being housed inside.  A peak under the hood reveals a ton of unused space (multiwell autosampler, perhaps!!!).  Pretty much every component on this instrument looks like a fraction of its counterpart on more common cytometers. However, it's quite clear that every penny possible was pinched in the manufacturing of this instrument.  Everything about it screams cheap.  That's not necessarily a bad thing per se, but as soon as you start opening up lids and doors and see some of the components inside, it becomes clear how they were able to create a functional instrument at bargain prices.  Beckman Coulter has said that part of what they will do to the CytoFLEX is to add some polish to the components without adding cost.  A final product and price point has yet to be revealed, but we expect to see it in the wild this fall.


The Sheath and Waste tanks sit beside the instrument and have a single output/input line, respectively. They hold about 5 liters, which should last most of a day with moderate use and reasonable amounts of backflushing. The preferred sheath for this system is some high quality H2O (Insert Waterboy reference here). Beckman Coulter will likely sell you a box of water at a premium and call it "Coulter Sheath" but you'll be just fine grabbing some DI from your MilliQ system.

Again the system moves fluid throughout using a pair of peristaltic pumps. The non-fluid movement of peristaltic pumps tend to make them not ideal for a system that requires stable fluid flow, but in testing the CytoFLEX, I saw no fluctuations in any of the channels over long runs with beads (plotting bead intensity vs. time). This type of instability due to peristaltic pump oscillation had been reported in some iterations of the Accuri C6 when it first came out. In the CytoFLEX, special attention was paid to create a pulseless peristaltic pump, which definitely holds true in my testing.

Although the sample volume flow rate has a custom setting that allows it to go up to 240ul/min, in
Close-up of the sample tube loading arm.
my tests, I saw dramatic declines in scatter profiles and less obvious, but still present, losses in resolution of fluorescence profiles beyond 100ul/min. I think the 240 setting would be great for cleaning the sample line out, or maybe forcing through a stubborn clog, but not for collecting data.  This type of flow rate is pretty much on par with other hydrodynamically focused fluidics system (unlike, for example the Attune that uses acoustic focusing and can easily go up to 1000ul/min with minimal degradation of profiles). Although the 80um wide beam spots may insulate the wide sample core stream from really poor resolution.

The sample loading stage is a bit funky at this point.  The loading stage moves in and out with the smoothness of 20 grit sandpaper sliding across berber carpeting (i.e. not smooth at all). This loading/unloading operation slows down the process just enough to be annoying, but you get use to it after a while. No plate loader (yet), No multi-tube loader (yet).


The optical system on the CytoFLEX is the biggest departure from any other instrument developed.  A lot of the technology is proprietary, and as much information as I was able to deduce I'll share here, but I could be flat wrong on some things, so take what I say with a grain of salt. 

Custom made Laser modules
Don't expect to see the familiar Coherent Laser Cubes on this instrument.  In fact, these lasers are custom made, in-house in the Chinese facilities (where the entire instrument is manufactured).  When you take off the laser compartment cover, you're greeted with non-descript tiny black boxes with a sticker on them telling you which laser it is.  Here is where they can save a lot of money.  Without being beholden to the Coherent behemouth, they're not locked into Coherent prices.  And, since they are making the lasers themselves, they can customize everything about them according to this specific instrument.  Worried about the quality?  I was too, until I saw the performance.  Of course, what I'm not able to test is long-term laser life on these.  The stated spec on Xitogen's web site for lifetime is 20,000 hours, but this hasn't been tested in the field, as far as I know.  The air launched beams go through the typical steering and shaping optics and terminate at the flow cell in front of one of a 7 "pinholes" on the instrument.  Beam sizes and power efflux are restricted to a 5um x 80um gaussian profile courtesy of the beam shaping optic and its large 1.3 NA, which means most of the laser power gets focused to the "pinhole" in a slit (N.B. I say "pinhole" since it's unclear if there are actual pinholes in the traditional sense or some other sort of voodoo magic).  This should allow for maximal excitation of fluors and minimal crosstalk between laser lines.
A look at the laser path with the covers off and interlock defeated
(Don't try this at home kids!)

Emitted light is collected by fiber optic bundles which carry the light to the detector blocks. The detector blocks, referred to as Fiber Array Photo Detector Modules or FAPDs is where all the innovation takes place.  The first thing you'll notice when looking inside the FAPDs is the small size of the filter sticks.  Pulling one out reveals a tiny piece of glass no more than a few millimeters square. However, the rest of the components inside the FAPD are completely foreign to someone who's looked at dichroics, bandpasses, and PMTs his whole cytometry life.  The light exiting the fiber passes through a wavelength division multiplexer, which acts like conventional dichroics to partially
Looking into the FAPD with a filter stick removed.
split the light into distinct ranges, and then the light is further refined by the bandpass filters before hitting the photodiode.  Photodiode? Don't you mean PMT?  No, you read that right, this system uses Avalanche Photodiodes (APDs).  These semiconductor detectors are well-known for their high sensitivity, and silicon based APDs have good quantum efficiency in the visible and near-IR range as well as low noise.  If they're so sensitive why haven't they been used before?  Good question, and as far as I can tell, the problem has always been the amount of voltage that needs to be applied to achieve high sensitivity and this high voltage causing breakdown of the APD.  Somehow, this has been circumvented in the CytoFLEX. The other interesting thing about the detectors is that the response of the APDs across the entire range is absolutely linear. They stand by this fact so much so that if you set up compensation on FITC vs. PE at one set of voltages, and then change the voltages, the system will automatically adjust the compensation values to take into account the new voltage settings.  This can only happen if the response is linear from end-to-end and therefore compensation is merely a mathematical equation with voltages as one of the variables.


The system uses 16-bit A/D converters and boasts of 7-decades of dynamic range.  Normally 16-bits doesn't get you that much range, but by oversampling the pulses at 40MHz, and adding up all the samples, a full 7-log scale can be achieved.  However, like most of these large scales, the first decade tends to exhibit poor resolution and is "hidden" by default.  So the scale goes from 10^2 up to 10^7. Qualitatively, I will say that I was able to resolve all 8-peaks of the 8-peak rainbow bead set with some room on both sides of peaks 1 and 8 - that doesn't always happen.

One of the only complaints I had about this instrument was the loss of data at moderate to high event rates. This has to be due to the pulse processing speed of the electronics system and its inability to process the signals fast enough.  It's likely influenced by sample concentration and the system's dynamic integration window - not unlike the FACSDiVa window extension setting.  If the window is reduced, % abort would likely decrease.  Also, increasing the threshold would also have the effect of better resolution between pulses and thereby decrease the abort rate. I did not explore either of these options when running and just used the default window extension and threshold. As you can see from the chart, even going at a moderate rate of 10,000 events per second yields an alarming abort rate. Going even faster results in a recovery of 50% or less. It's important to separate your ideas about % aborts on analyzers from high-speed sorters.  Cell sorters have the advantage of pushing the cells through at very high velocities resulting is narrower pulses and an easier time resolving two closely related pulses.  But, on analyzers, the cell velocity is much slower, resulting is broad pulses and more difficulty resolving closely related pulses. Therefore, the abort rates are typically going to be higher on slow flow analyzers, however we're not as aware of these abort rates on analyzers because we are
always only concerned with frequencies of populations and not absolute yield of populations (like on cell sorters).  So, 10% abort over 20,000 events per second might be reasonable, however, 10% abort rates over 10,000 eps is probably not.

For this test, I created a concentrated sample using a suspension cell line, which, at 60ul/min should yield 50,000 events per second.  I then created serial dilutions from there all the way down until an expected 2,500 events per second.  I ran each tube on the instrument and recorded the event rate displayed by the system's counters. 


If you've used CellQuest and FACSDiVa in your cytometry lifetime, you'll feel right at home here. There's not much to say about the software except that it works.  It was super easy for me to pick up.  I was shown nothing as far as how to operate the instrument, do compensation, etc... and I was able to figure it out with minimal struggle. The CytExpert software does one thing really well and that is it gives you a large, unobstructed view of your data, and just enough controls in a thin side panel to allow you to acquire data.  I'm sure there are some analysis tools built in, but I don't care, I just want to acquire data, dump it into FlowJo and worry about analysis later. It gives you the ability to do automated compensation, use biexponential display, perform gating, and show stats windows. It would be interesting to see if something like Kaluza-G would ever make it onto something like this. But then again, Beckman Coulter already has 900 acquisition softwares already, what's one more!


Finally the good stuff. What can I say, this thing rocks. In terms of fluorescence sensitivity, it beat the pants off of anything I've ever tested full stop. I've put a range of values together for fluorescence resolution that shows the spread of instruments I've tested.  The value (called qNORM) represents the lowest number of antibodies bound that can be resolved from unstained lymphocytes.  The lower the value, the better, and as you can see, the CytoFLEX, with its APD detectors and DIY lasers easily beats the average across the board.  Of course I ran all the other common bead sets on this instrument.  Everything I threw at it, it handled with ease. 

8-Peak Resolution at low and high flow rates:

As you can see, resolving 8-peak beads is a cinch on the CytoFLEX pretty much across all channels. Even at the highest flow rate (240ul/min) the fluorescence resolution remains relatively unchanged, however the light scatter experiences some funky spread at the high flow rate.

APD Voltage Optimization:

Using a blank bead, the voltage was moved up and down the scale at appropriate intervals.  The rCV was calculated on the single bead peak in each of the fluorescence channels. Using a similar test as PMT optimization, I wanted to see if the APDs behaved in a similar way. It does appear that there is a sweet spot for APD voltages that vary across parameters.  This mimics PMT optimization profiles commonly seen before.


A pretty simple linearity test using PI stained CENs, and everything checks out as expected.  However, like I mentioned earlier, linearity on this instrument has a bigger role on this instrument than others.  With the CytExpert software, you can setup compensation at one set of voltages, change voltages (because it's a different cell line, or the sample is too bright, or other reason), and it will recalculate compensation based on the new voltages.  This may have been (or may still be) part of the FACSVerse software, but I've never used one of those, so I'm not sure about that.  Theoretically, then, you could create a set of comp tubes using non-tandem antibodies once, and then recall those comps each time, even if you've changed voltages. Anywho, linearity is great, it deviates from the theoretical line by less than 1% across the board.
PI Stained CENs comparing the theoretical line and the actual data

qNORM Resolution Comparison:

Without going to much into the methodology (because I've done it so many times before), what follows is a comparison of pretty much every instrument I've ever tested (grey boxes with quartile whiskers) with the CytoFLEX (blue circles) overlain.  This metric measures the number of bound antibodies that can be resolved from unstained lymphocytes. So, lower numbers equals better low-end resolution.  As you can see the CytoFLEX compares very well with all the best instruments out there. It definitely beats every instrument I own in the FITC, PE, PECy7, and APC channels.  In the PacBlue channel, it's about average.  I'm pretty sure this system can resolve pretty much any dim population you can throw at it.

Final Thoughts:

It's evident to me that the CytoFLEX would suit the needs of many demanding applications.  There's really no questioning its performance in terms of fluorescence detection. Light scatter resolution of cell populations wasn't as good as some of my better instruments. However, small particle detection, especially using 405nm side scatter is reported to give <200nm resolution.  I typically don't test small particle stuff since it's not really my thing. The fluidics seemed stable and robust for the time I had it.  I ran as many cell samples as I could to see if I could clog it up or make things look bad, and other than the high abort rates at high event rates, I saw no issues from a fluidics standpoint.  The software is fine for what it needs to do.  I'm so entrenched in doing analysis in FlowJo that I couldn't care less if there are histogram overlays or other fancy analysis-only plots in my acquisition software.  I just want it to be fast and simple to use. 

We also don't really know about the long term reliability of the hardware components. Sure everything works fine over the course of a month, but what about a couple years or more.  Will it have the staying power and uptime of a FACScan? This, I'm afraid, only time will tell.

But, I think the most important take home message here is that this instrument proves that flow cytometry hardware is absolutely a commodity in the eye of the consumer. As fancy as one wants to make hardware these days, no one is going to be impressed.  And the fact that hardware can be made cheaply reinforces this fact.  What this means is that we'll finally see a shift in focus away from over-engineered hardware to hardware that just works, but this time with a super slick user interface that people are attracted to.  The future is all about software and services, and I, for one, couldn't be happier!

Postscript: At the time of publishing, Beckman Coulter launched a new splash page with specs and a glimpse of the new exterior of the CytoFLEX.  You can reach that page here

Wednesday, January 29, 2014

21st Century Learning using an Ancient Model Applied to Flow Cytometry Training

There are so many things I'd love to learn.  I always imagined myself playing the guitar, or executing a no-hands backflip, or even writing a mobile application, but so far, I still cannot do any of those things. Of course, each of these things are certainly in the realm of possibilities for me.  I'm actually somewhat musically inclined, I can do many gymnastic-type flips, and I know a thing or two about the languages of the coding world, and yet, I can't amend my CV with any of these goals.  So, how might I go about learning these skills that envelope both knowledge of abstract concepts (like coding) and physical moves (like playing a guitar)?

I don't know about you, but whenever I'm trying to figure something out, I default to YouTube. YouTube is great for things like this, in that you can pretty much find a video demonstrating something on any topic you're interested in.  However, where things tend to fall apart is the one-way nature of YouTube.  The demonstrator is broadcasting out a message that I may stumble upon years after it was uploaded and there's not a great way for me to interact with the original creator.  Sure, I could leave a comment in the hopes that it'll be answered, but I'm just as likely to get an unhelpful snide remark.

In fact, many models of learning these days follow a similar strategy.  E-Learning is all the craze these days.  Popularized by online e-learning houses such as Kahn Academy or Coursera or even on a larger scale, institutions such as University of Phoenix, e-learning promises all the bang for little to no buck, all taking place in the confines of your oversized easy-chair. But, questions remain as to how effective these programs are. I know I've signed up for courses a few times only to stop going after the second or third lecture. In some ways the information is presented as little more than a canned powerpoint presentation with some voice-over description.  A step up from here is the possibility to interact, real-time with the presenter via chat or video conference.  Even, with the best implementation of these technologies, remote e-learning is difficult.

Let's flip this conversation completely on its head for a moment.  For millennia, the way in which people learned a trade or skill or gained any sort of knowledge was through a Master/Apprentice process.  The elders of the group who had the necessary experience and expertise took a young apprentice under his wing and taught him the way.  If it helps, I always conjure up images of Qui-Gon Jinn teaching his Padawan Obi-Wan Kenobi in the ways of the force.  You could imagine the education the apprentice received was really good, but the process was somewhat inefficient since a Master may only have a limited number of apprentices. Contrast this with e-learning and the dichotomy should be clear.  E-learning may provide a highly efficient means of disseminating information, but the actual learning may be inadequate whereas the Master/Apprentice model may provide for world-class learning but is inefficient in terms of disseminating information to a large group of eager learners.

To bring this conversation closer to home, how do we go about teaching the art of flow cytometry to the next generation of scientists?  I would say, up until this point, the passing on of flow cytometry knowledge has favored the Master/Apprentice model.  This is certainly the way I learned, and probably the way I would prefer to learn just about anything.  In recent years, however, many core facilities, companies, and professional organizations have tested out the e-learning model of teaching flow cytometry.  Like the e-learning trailblazers, these differ in quality from powerpoint slideshows to interactive, well-produced, and highly animated videos.  We've tossed around the idea of moving towards an e-learning model at UCFlow, and what's held us back (aside from the technical complexities involved in producing something worth putting your brand on) is a core belief that we can produce better cytometrists with the more intimate master/apprentice model.

The question then becomes, can we leverage modern communications technologies to make the master/apprentice model work more efficiently?  Well, of course the answer is yes, otherwise I wouldn't have bothered writing this post.  But, instead of describing a fictional method in overly verbose prose, I want to point you to what I think is the ultimate model of learning PERIOD.

It pairs the master/apprentice model with new technologies like video conferencing, chat, hangouts, google glass, wearable tech, etc...  It also gamifies the process to promote better engagement.  Imagine this scenario.  I know a little flow, but I'm faced with this new application.  I'd really like to start doing microparticle analysis.  I log into the cytometry masters portal, and search microparticles.  Up pops a list of microparticle experts with various specialities and levels.  For example Jane is a level 50 Endothelial MicroParticle Master, and can accept a new apprentice for the next month.  She prefers to communicate via Google+ Hangouts and is in the Pacific Time Zone.  I connect with Jane, learn all her tricks and tips, and then I can level-up in my knowledge of microparticle detection, bringing me to a level 10 master.

Would you like to see how this works?  Luckily, this has already been launched using a different, but I'd dare to say very similar, technology - photography.  The super awesome photographer, Trey Ratcliff ( launched a brand new site, The Arcanum ( that uses this exact model.  Master/Apprentice, Modern Communications Technologies, Gamification.  Watch the video below to see what it's all about.  What I love about this is that it's visual; you're learning directly from an expert of whom you can ask all the nuanced questions you like; it uses all the latest gadgetry; and the gamification of the process makes it way more engaging.

The next question I have is, Who wants to build the flow cytometry version of this with me????

Monday, November 18, 2013

He said, She said, PEBCAC?

In flow cytometry core facilities, scenarios such as the one that follows are commonplace.
The first thought that comes to me
after a user reports a problem.
An end-user is attempting to collect data, for some reason there's an issue, the end-user requests assistance from the core facility staff, some resolution is achieved, lather, rinse, repeat.
But, the interesting thing is the back and forth between facility personnel and the user.  Each party is trying to figure out in what way the other party messed up the experiment.  A veritable "he said/she said" ensues and eventually a resolution is achieved.  The way in which the resolution comes about can take many forms depending on the level-headedness of the parties involved. However, core facility personnel are typically about as protective of their instruments and services as a mama grizzly is towards her newborn cubs.  Similarly, a precocious grad student, who has spent umpteen hours preparing her samples, couldn't imagine a situation where she could have made an error.  To celebrate this perennial back and forth, I present to you the 10 most common phrases (5 from each side) overheard between core facility personnel and end-users during the initial throws of an experimental/instrument mishap.

5 Most common statements from core facility personnel when presented with a problem by an end-user

  1. Did you try and reboot the instrument (software)?
  2. Hmph, my QC beads look fine.
  3. Did you filter your samples before bringing them here?
  4. I don't know... everything looks pretty dead/negative to me.
  5. No one else has had any problems on here today.
5 Most common statements from an end user when they encounter a problem at the core facility

  1. Why does this thing break every time I try and use it?
  2. I had X million cells, so why did the instrument only run X/5 cells?
  3. The instrument is clogged or something.  The person before me didn't clean it well enough.
  4. Well, will the problem be fixed soon? This data is for a grant proposal due tomorrow.
  5. I hope you're not going to charge me for this.
Of course, I'm a bit biased when it comes to this scenario, so you may have your own favorite anecdotes to share.  You can do so in the comments. Flame on!

Tuesday, July 23, 2013

10 Tips for purchasing your next cytometer

So you've got some money to spend and you figure, heck, I do so much flow, maybe I'll just buy a(nother) cytometer.  Presented here are some tips to avoid buyer's remorse, see though the marketing spin, and make an educated decision on which instrument to purchase.  But, before we get into that, let me first state that I'm NOT going to make this decision for you and tell you which instrument to buy.  I am merely going to provide you with the tools to make as good a decision as you can.  In fact, these are the very same steps I go through whenever I'm in the market.  And so, I present to you, 10 tips for purchasing your next cytometer.

#1.  Define your needs.  What are the applications you will run on this system?  How many parameters (realistically) do you run on average?  How many parameters will you run in the near-future?  Are there any specialty dyes you run?  Do you prepare samples in tubes or plates?  At what event rate do you run your samples?

Example:  I have some projects in mind which require 8 - 10 fluorescence parameters.  At 3 parameters per laser, I probably need a 3-laser system minimally.  I like to stain/run my cells in a 96 well plate, so a plate sampler option is needed.  My experiments typically involve immunophenotyping rare subsets, so I collect 10^6 cells at rates of about 15,000/second.  I don't need to sort.

#2.  Query the end-users.  If others in the lab or core facility will use the instrument, ask them the same questions as in #1.

Example:  Another user in the lab does a lot of screening of her mCherry transfected cell lines.  She doesn't collect a lot of cells, but screens many samples.  She would require a yellow/green laser for excitation, and fast 96-well sampling capabilities.

#3.  Refine your needs.  Combining the information you learn from steps 1 and 2, you should be able to refine the needs for this instrument.  Annotating this list, and possibly triaging needs and wants will be very helpful at this point.  Of course you are working within a budget, so you'll certainly want to keep that in mind as you survey the market.

#4.  Survey the market.  You probably already have an idea of the "big" players in the market, but even if you didn't, simply typing the query "flow cytometer" in your browser brings up 9 different instrument manufacturers within the first two pages of a Google search.  You can follow these links, collecting information on the various instruments.  For something like this, I like to use an electronic note taking application like Evernote to keep everything together, and make notes as I go through the process.  For those web sites where information and materials are not easily accessible, sending an email to a local sales representative will get you all the marketing materials you could ever ask for.

#5.  Learn how to read marketing materials.  Speaking of marketing materials, there are a few things you should be aware of.  Beyond the very basics (lasers available, number of detectors, etc...), much of what you see in your average cytometer specification sheet is useless information.  It's basically a list of values for meaningless metrics that MUST be put into the materials to match what the competition is stating.  For example, the ever-present detection threshold of FITC and PE.  Most all spec sheets will include something like an MESF Detection Threshold of <150 for FITC and <100 for PE.  This means absolutely nothing in terms of how well the system will work for your applications let alone how other colors will fair.  You'll also see outrageous specifications for event rates, like 100,000 events per second!  Lastly, and probably my favorite, is the panel of histograms showing the resolution of multi-intensity hard-dyed beads (e.g. 8-peak Spherotech beads).  You can pretty much ignore all this information, and focus on the things that matter.  How many lasers?  How many detectors?  Can you upgrade the system in the field with more lasers/detectors? Is there a multi-well sampler? etc...

#6.  Create the matrix.  By now, you have a list of needs/wants, and you have a bunch of marketing materials.  Put it all together in tabular format.

Fictitious Instrument Comparison Chart, with the 3 contenders.

#7.  Gain hands-on experience for the top 3 contenders - make sure the OEM knows the fate of the sale hinges on the success of the hands-on demo. Run your battery of tests that matter to you, evaluating the results, as well as ease-of-use, software, UI, hardware.  Simply staining your favorite panel of antibodies and running it on the instrument will give you a TON of information as to how these cytometers stack up.  Running real samples (not just beads) is an absolute must.

#8.  Take to the social network (and take everyone's opinion with a grain of salt).  Useful things that can come back from the community include; recurring hardware/software failures, maintenance issues, service issues, responsiveness, etc...  For any negative responses that come up, make sure to bring these to the attention of the manufacturer (respecting people's confidentiality, of course) and ask for a response.  Get everything in writing.  No phone conversations!

#9.  Negotiate the purchasing terms with multiple companies.  Make sure the sales representative is aware of their competition.  Aside from asking for the best possible price, discuss other value added options.  For example, an extension of the warranty, free training slots, discounted multi-year service agreement, free shipping, free upgrades (higher powered lasers, multiwell samplers, extra emission filters, next version of software).  Get everything in writing, no phone conversations (did I mention that already)!

#10.  Take advantage of year-end discounts.  If possible, time your negotiations and purchase with the end of the company's fiscal year.  You'd be surprised what sort of deal you can get if the company is close to reaching their target for the year.

BONUS Tip:  Don't be afraid of venturing away from the "big companies."  When dealing with newer companies and newer technologies, getting cutting-edge hardware can be a double-edged sword. Although you may be able to get a deal on price, make sure there are some protections in place that allow you to get future upgrades or revisions to problematic hardware for free.  At the very least, you should be able to get a percentage of your money back if it's a total failure.  Again, get it in writing up front.

So there you have it.  I think if you keep these common sense tips in mind when purchasing your next cytometer you won't be disappointed.  Got any other tips that have helped you make the right purchasing decision?  Why not leave a comment below.