The most sensitive Cytometer available?

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

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.

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

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?

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"

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.

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.