Compensation is infiltrating my dreams.

I've been composing this post in my head for a couple of weeks now, but have been too busy to sit down and write it, so it shouldn't really surprise me that it popped up in a dream.  However, a new twist was added via my unconscious mind (which I'll get to later).  So, the original post was all about how I've pretty much given up on compensating using cells, and if you're not using beads, then you're pretty much setting yourself up for compensation failure (unless of course you're using things like PI, or mCherry, or the like).  I mean, the whole point of 'autocomp' is to take the subjectivity out of compensation, and using objective mathematics to correct for fluorescence spillover.  However, every single time I've done autocomp using cells, it just doesn't look 'right' and I end up tweaking the values just a little bit.  I've come to terms with this fact, and have pretty much settled with this sub-par situation.  But, if you're trying to teach someone about compensation, and you introduce this 'autocomp' feature, it makes for a pretty awkward conversation when you then go on to say, "Well, just adjust the values a little bit until it looks right."  So, I typically recommend people do their compensation with beads.  For many of my users, the thing that prevents them from doing this is cost, or maybe a bit of skepticism in changing the ways they were taught to do their staining.  The reasons why compensating using cells doesn't always work are many, but let me just outline a few for you here.

1.  Insufficient frequencies of both positive and negative fraction to make a statistically significant regression of means.  If in your stained cell sample, you only have a 0.1% positive fraction, the mean of that population in the spillover channel will not reach a high enough statistical significance until you collect millions of cells.  No one is going to collect millions of cells on their single stain control.  This also holds true when all your cells are positive for your single stain control, and you have a really low negative (or low) population.

2.  Poor resolution of the positive fraction.  Sometimes you will not have a clear positive population, so making a gate around the positive fraction for performing compensation is difficult.  If you end up encircling some of the high autofluorescent cells that you mistakenly call positive, your compensation will surely be off.

3.  Non-linearities at the extremes can lead to inaccurate compensation.  If you're compensating using an unstained (or negative) fraction that is at the very low-end of the scale, or if your positive fraction is at the very high-end of the scale, you're likely using a data point in the non-linear range of the log scale. Since compensation algorithms are basically relying on the fact that your range of analysis is linear, you're going to run into lots of problems if you're using "unstained" cells as your low-data point, or really bright cells as your high data point.  Side bar:  Yes, I know, your comp control should be at least as bright as your sample staining, blah, blah, blah.  However, the only reason why this is the case is because of non-linearities at the very high end of the scale.  If all your staining fell within the linear portion of the scale (let's say 1.0 logs to 3.5 logs), then this isn't necessarily a problem.  You can take any two points within that range, and create a regression line that will model the entire scale.  No-scale is linear enough, especially at the extremes, so the 'rule' of a maximally bright comp control needs to be adhered to.

4.  Mismatched autofluorescence between positive and negative.  If I stained my leukocyte prep with a monocyte marker (CD14, for example).  All my monocytes will be positive.  For this single stained comp control, what should I use as my negative?  Many people would simply use the negative lymphocytes or granulocytes, and many people would end up with a poor compensation matrix.  For channels where autofluorescence is a factor (mostly the green/yellow detectors off the blue and lower laser lines), the positive fraction's autofluorescence should match the negative fraction's autofluorescence.  This is, evidently only necessary when you're using cells for compensation, and you have a mixed cell-type sample.

So, there are certainly lots of pitfalls when using cells for compensation, which is why using beads is a good idea.  To solve many of these issues, simply using an antibody capture bead at two fluorescence levels should do the trick.  You'll notice I said two fluorescence levels, and not one positive and the 'blank' bead.  Using the blank bead can lead us into issue #3 above, so I prefer to use the bead at a saturating level of antibody and maybe 100-fold less, to create a high and low peak.  In the end the peaks will fall around the 3.5 decade range and 1.5 decade range.  Use these peaks as your 'positive' and 'negative' values in your favorite autocomp program, and voila, perfect compensation.  Of course, these beads are run at the appropriate voltage that is set up according to your cell type.

But, what about the twist?  The twist is, that you don't need to only use beads as your capture matrix.  You could use cells!  I know, I know, I just went on and on about NOT using cells, now I'm telling you to use cells, but wait, let me explain.  Take a thymus, get all your non-tandem antibodies in CD4, stain them at two concentrations, fix them, and stick them in the fridge.  You now have ready-made compensation controls that are much cheaper than buying capture beads.  Why thymus?  They're the closest thing to beads; pretty much homogeneous, so we don't have to worry about autofluorescence mismatch, they're almost all CD4 positive, so that makes it easier to create two nice peaks, and you can get a boatload of them from a young mouse.  On top of all this, we gain the ability to use other things besides antibodies.  You could stain them with many of your dyes for a comp control, PI, DAPI, CFSE, etc...  Something you can't do with beads.  For tandems, I'd stick with capture beads.

Ok, there you have it.  If you've made it this far reading through all my gibberish, let me know what you think.

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

Ah, the MoFlo - what a fine piece of craftsmanship!  I started my relationship with the MoFlo (Formerly of Cytomation, Formerly of DakoCytomation, Formerly of Dako, Currently of Beckman-Coulter) in the year 2000.  We had many great years together, but our relationship was getting a bit stale.  You see, there was this fancy new gal in town call the Aria who lured me into her web of seduction with promises of 'turn-key' operation, and I bit!  I soon realized however, that the grass isn't necessarily greener on the other side, and re-visited the rock-solid usability of the MoFlo.  In recent years, the MoFlo started showing its age.  I have to admit, part of the issue was a certain level of neglect and abuse on our part, but hey 10 years in instrument years is like 80 in people years.  And so we came to a fork in the road, and as with most things in the technology area utilizing 20 year old components, we had to decide, pull the plug or pursue the upgrade path.
When I was contacted by the folks at Propel labs (who, evidently are a group of people from the original Cytomation company) that there was an upgrade path to the XDP electronics for the legacy MoFlo, I was thrilled.  After about a year of begging for money from anyone that would listen to me, I finally secured the funding and was ready for the upgrade.  So, why upgrade to XDP instead of buying a new sorter?  Well, first of all, it was a financial thing.  The cost of an upgrade is about 1/4th the cost of a new sorter.  Secondly, the fluidics on our MoFlo are uncannily stable; who knows if we'd strike it rich again with a new sorter.  You may also be asking, what's so great about XDP?  Well, I'd never be able to explain with such elegance as Dan Fox could, so all I can say is track down the white paper Dan wrote, read it, then pick your lower jaw up off the floor.  The big lure for me (besides the obsolescence of parts for the legacy MoFlo) was just the fact that we'd be able to operate with no/low hard aborts similar to the Aria, which, when paired with the higher number of droplets a jet-in-air can achieve, should allow us to sort faster and maintain high yields and purity. With our XDP upgrade, we also had all our PMTs changed, and threw on two new laser lines to boot.  - Side note - We had one of these co-lase towers installed on our MoFlo, which is also a product of Propel Labs, that basically combines two laser lines so they can be run colinear into the 3-pinhold MoFlo setup.  We chose to put on a UV and Red laser and run them colinear through the co-lase tower.  This now gives us a 4-laser MoFlo (355, 488, 561, and 640) - End Side Note -

The remains of the MoFlo after the tear-down

As far as the actual upgrade goes, the install went pretty smooth.  It took 2-3 guys about 3 days to completely tear down the instrument to basically an empty table, and then install the PMTs, electronics, the touch-screen panel, and the computer.  As with most installs/upgrades, we did have a couple hiccups, but they were taken care of immediately.  I guess that's one good thing about working with a smaller company like Propel Labs.  They can't afford to lose any business, so customer service is automatically very good.

We've been using the XDP now for about a week, and things have gone pretty well.  We're still getting use to the touch-screen interface, and some of the new things in Summit, but overall, I'd say we made the right decision, and hopefully the MoFlo can dutifully give us another 10 years of service.

Once we've gotten into a rhythm on this thing and really test the bounds of speed, I'll post some data.  But for now, enjoy a pic of the finished product below.

The upgraded MoFlo XDP in all its polished glory!

Follow us on Facebook!!

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

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

GLIIFCA Core Manager Meeting Preview - September 24, 2010

For those who will be at the meeting, I've put up my slides in PDF format here (sorry, I think the link works now) in case you wanted to follow-up with one of the tools we use in the core.  For those who will not be able to attend, feel free to read through to see what types of tools we use at UCFlow to try to do more with less and be as efficient as possible.  Check back here for updates from the Core Manager meeting and the rest of this year's GLIIFCA.

Flow cytometry leads to everything!

The man on the right in this picture is Jeff Schneider.  He used to be a technician here in the flow lab.  Look at him now, playing with his band Darling at the Hideout tonight at 9PM.  Rock on dude!

Flow cytometry leads to everything.

You know what really grinds my gears?

So, I was fixing a clogged DCM pump on the LSRII this morning, which requires the removal of the side panel on the instrument, getting on your hands and knees and digging around in the inner bowels of the beast.  As I was shimmying around, contorting my limbs in all sorts of god-awful positions, I kept crunching pieces of plastic under my feet.  I peaked down, backwards, over my shoulder to see a bunch of pipette tips on the floor...surrounding the trash bin at the foot of the instrument. Are you kidding me?  People, presumedly adding their PI or DAPI at the instrument, are ejecting their used tips in the vicinity of the garbage, and missing >50% of the time.  They obviously can hear the tips crashing into the ground and missing the garbage, but decide to do nothing about this?  I then pan across the room, and I see racks of nearly empty tubes, crumpled kimwipes and a full waste tank.  C'mon people, have a little respect, pick up after yourselves.   Needless to say, my DCM repair turned into a full-on cleaning session; swept, threw out all the tubes, wiped down the bench space, and even got the mop from the janitor's closet and mopped the floors.  So, here we go people, August 27th, 2010 at 10AM, the LSRII area in 037 is clean.  LET'S KEEP IT THAT WAY!

It's GLIIFCA Time!

We're in the dog days of Summer, so that could only mean one thing... GLIIFCA 19 is right around the corner.  The Great Lakes International (because we let the Canadians join us) Imaging and Flow Cytometry Association's 19th annual meeting will be held September 24 - 26, 2010 in Detroit, MI.  This is a great meeting for users of Flow Cytometry and Imaging technologies (including the ImageStream!) that gives you a chance to see how people are using Flow in interesting and novel ways in their research.  It's also ridiculously cheap.  The registration fee is a paltry $80 AND, if you bring a poster, you'll get a $100 travel stipend.  Also, it's close enough to drive,  and the hotel rates are very reasonable as well.  So, there's really no excuse not to go.  However, if you need one more reason why you should go to GLIIFCA this year, it's because the theme for this year's Saturday night party is freaking STAR WARS!!!!   So, click on over to GLIIFCA.org and register yourself today.  If you U of C folks do decide to join us, David will buy you a beer!