Drop-seq Tutorial & Troubleshooting

***Please note that we are not currently hosting Drop-seq tutorials. When future tutorial sessions are available, an announcement will be posted both here and on our google forum.***

Hi everyone!  While nothing can beat hands-on experience and we continue to welcome fellow Drop-seqers to visit us for an in-person tutorial session, I thought it would be helpful for there to be an online version for those who can’t make it to Boston (and also to serve as a source for people to refer back to).  Just as we have created a living protocol, this page will continue to evolve over time, so feel free to reach out to me with additional questions or video requests.

If you would like to visit us in Boston for a tutorial session, send an e-mail to dropseq@gmail.com with the subject line “DROPSEQ TUTORIAL” and provide the following information:

  1. Your name
  2. Your lab & institution
  3. How many people will be attending
  4. A range of possible dates


Alright Drop-seqers, let’s get started!


In our paper, we recommend a cell occupancy of 0.05, or 5%.  In 125 um droplets, we load our cells at 100 cells/ul because this results in a final concentration of 50 cells/ul within our droplets, since half of each droplet’s aqueous volume comes from the cells, and half of the volume comes from the beads.


So say that you have devices that are producing droplets that are 100 um instead of 125 um.  Smaller droplets are perfectly fine to use, you’ll just want to increase your loading concentrations accordingly.  For a droplet of 100 um, its radius is 50 um.  Therefore, the volume of this droplet with a radius of 50 um = (4/3)*pi*(50^3) = 523,000 cubic microns = 0.523 nL.  Since the volume of a 125 um droplet is 1 nL, you want to take the ratio of the volumes and then multiply by the standard loading concentration:  (1 nL / 0.523 nL) * 100 cells/uL = 191 cells/uL.


Because our current devices produce smaller droplets, we use a higher loading concentration for our cells and beads than is published in our paper.  With a cell loading concentration of 220 cells/uL and a bead loading concentration of ~260 beads/uL, we typically expect to recover around 6K-7K STAMPS per mL of cells flowed through a device and collected in a single falcon tube.  Note that the number of STAMPS you recover will depend on how clean your break is, and how many beads you lose during processing.




I’ve been getting a lot of questions lately regarding how to set up the V&P mixing system.  Here is a photo of the latest Drop-seq system that we set up in another lab.  I recommend always purchasing the V&P accessory kit (VP 710D2-4), then use your imagination to get the angles right!





Counting beads

Because the beads are so dense, it is important to use a P200 pipette set to 20ul and not a P20 or P100, since these do not have enough force to pipette the beads fast enough to prevent their settling inside the tip.  Preload your tip and have your C-chip positioned next to it, mix up your beads with a P1000 or pipette aid, and quickly use the P200 to suck up beads from ~the middle of the suspension and transfer it into the C-chip in a nice fluid motion.  When you look at the C-chip by eye, you should see the white dots (beads) evenly dispersed.  If you see a white streak down the center or the beads don’t look uniformly dispersed throughout the C-chip, your count will not be accurate.  Practice makes perfect!


Here’s what evenly distributed beads should look like in your C-chip at a concentration ~280 beads/ul.

Evenly distributed beads




















Bead clogs and “vanishing” beads

Loading a bead concentration greater than 300 beads/ul significantly increases the likelihood of a clog.  While it is always good practice to record your bead concentration, your bead count is not as vital as your cell count because when your bead concentration increases, the number of droplets that contain a bead increases, but so does the number of “empty beads” – droplets that contain a bead but no cell.  Therefore, the proportion of beads that capture a cell remains the same, although the number of bead doublets will increase.

There are a few key practices that help prevent bead clogs when you’re setting up for a Drop.

  1. Always use a 3ml syringe, never use a larger syringe for your beads.
  2. Never load more than ~1.5ml of beads into the syringe – the magnet cannot sufficiently mix a larger volume.  We typically stick to 1.2ml.
  3. Make sure you mix your beads before putting on the needle, then keep it mixing uniformly throughout the experiment (the disk should be moving up and down through the full volume of your beads).  If you fail to do this even for half a minute, the beads can settle and either become clogged in the needle tip, or they will flow through the device at a significantly higher concentration at the beginning of your run and by the end of your run you’ll see that there are hardly any flowing through.


Good mixing

Note how the magnet is traveling vertically through the full volume of the loaded beads.  The angle of the magnet is much more important than the exact distance.




Bad mixing

Note that the beads look uniformly suspended because I only did this for a few seconds to take this video – over time the beads would have settled toward the bottom of the syringe.  Of course as your volume of remaining beads decreases, eventually vertical mixing is no longer possible – at which point you should adjust the magnet accordingly.



Under the scope

What magnification should I use and what should I be looking at?

You always want to be looking at the device on 4X magnification.  Having it zoomed in farther might create the illusion of what we call the “clear road” after the triangle, and also prevents you from seeing the full picture.  When monitoring a run, you need to be able to simultaneously see the bead inflow chamber (to ensure that beads are flowing in and aren’t clogging where the chamber narrows, which is the most common place for a clog), the triangle, and the blurry trail of droplets to the right of the triangle.  Sometimes you’ll see your cells going in, sometimes you won’t.  Due to small back flows of oil (which aren’t a problem), you will notice that the bead and cell inflow chambers will at times be partitioned off in different ways, which in turn means that the beads or cells will all be shooting through a smaller opening – this is why sometimes you might see your cells, and other times not.  Note that sometimes the oil can appear black as in the first image below, and other times you may just see large clear bubbles like you see in the second picture.  (The bubbles should stay in place or perhaps eventually clear out in the span of a few seconds – if you are seeing continuous bubbles going into the device that is very bad).  The most important thing to remember is that if your triangle looks right, everything is flowing in as it should be.

Oil but good drop













































Bubbles but okay



































So what is the “triangle”?

The triangle is the junction where all the flows (beads, cells, and oil) come together to form the droplets.  You want to play with the focus until you can clearly see the outline of the triangle.  Knowing what a good triangle looks like is key to performing a Drop-seq experiment, because it a) shows you that you’re producing uniform droplets, and b) allows you to troubleshoot any flow problems that might be contributing to poor droplet quality.


Examples of a good triangle

A “good” triangle does not always look exactly the same from one device to another.  Sometimes it will be slightly wider, or narrower, or longer, or shorter.  The shape of the triangle is impacted somewhat by the flow rates you’re using, which may differ depending on the quality of that particular device.  However, you will learn that while minor changes to shape can still generate great droplets, more drastic changes are your first sign that something is wrong.  If your triangle does not look right, your droplets will not look right either.  Only check the droplet quality using the run-down method, rock-method, or under the scope on a C-chip once you have achieved a stable triangle.

Good triangle 2

Good triangle 1















Good triangle 3
















Here’s a triangle that’s a bit wider at the base, but it still produced uniform droplets.  When your triangle seems a bit on the skinny or wide side, it’s always good to check (by eye) that all tubing is properly affixed to the needles and that there is no leaking either by the needles or where the tubing enters the device.  You should also look at a small aliquot of droplets in a C-chip toward the beginning of your run to make sure that quality is high (I typically only do this when the triangle looks a little funky).

Good triangle





















When your cell flow decreases  the base of the triangle widens and then curves outward

This could be either because a) you’re out of sample and your run is finished, or b) there is something wrong (cell solution leaking through a nick in the tubing, needle not affixed properly to the syringe, pump error, etc.)

Cells out

Cells out close-up


























When your bead flow decreases  the triangle becomes very narrow and gradually elongates

This could be either because a) you’re out of beads and your run is finished, or b) there is something wrong (bead solution leaking through a nick in the tubing, needle not affixed properly to the syringe, pump error, beads clogged in needle or tubing, etc.)

Beads out

Beads out close-up


























When your triangle is pulsing

There’s no missing this – sometimes the entire triangle will pulse like a heartbeat, in and out, in and out.  If you see this, you are producing droplets, but they will be dramatically different in size.  When you see this, it usually just means that your magnet is either touching the bead syringe or pump, or sometimes (though more rarely) this will happen because it is a bit too close to it even though it isn’t touching.  Since getting your magnet in the correct position is key to good mixing, the easiest solution is to simply slide the bead syringe over, as shown below.



When the tip of your triangle is flickering

When monitoring a stable Drop-seq run at 4X magnification, the triangle should appear almost unmoving.  If the end of the triangle (to the right) is flickering or jumping around, it means that you are not producing uniform droplets.  There are two approaches to fixing this problem.

  1. Decrease the flow rates of the beads and cells.  Try decreasing both by a few hundred ul/hr and see if the flickering improves.  Remember, always slow down the beads before the cells, and if you decide to increase the flow rates, always increase the cells before the beads.
  2. Try taping the outflow tube down to the microscope stage to stabilize it, and make sure your outflow tubing isn’t longer than necessary – it should reach the bottom of your collection tube, but if it’s much longer than this it could be contributing to instability at the triangle.


Debris in the device

So long as your triangle looks good and droplets aren’t being sheared between the triangle and the outflow hole, debris shouldn’t make a difference.  The image below shows some debris in the top cell channel and directly to the left of the triangle (this is just dust that got introduced during production), as well as dust that was flowed in with the beads that is trapped right where the beads focus into the channel that forms the triangle (looks like a strand of hair).  There is also a smaller bit of debris in the bottom oil channel.  When you see this hair-like debris in the bead chamber be alert, because a bead clog could occur at any moment.  However so long as the beads are continuing to flow through (even if several do get stuck to the piece of debris), you should continue collecting.

Remember- if your triangle’s okay, you’re okay!

Debris and hair




































Sometimes a piece of debris will get lodged after the triangle in what we call the “elbow” and shear the droplets – this is impossible to miss, since the blurry line will turn into a “clear road” after the piece of debris.  Below is a piece of that hair-like debris stuck in the elbow, but since the outflow remains blurry after encountering it, it isn’t shearing the droplets and is perfectly fine to leave as is.

Debris in elbow
































When you accidentally put the oil tubing in the cell inlet, and the cell tubing in the oil inlet

Often you will still be making droplets when this happens (you will see a blurry road), but the shape of the triangle is entirely off!  Here’s a sketch of what shape you would see, followed by a few droplets emerging to the right:

Screen Shot 2016-05-09 at 5.56.43 PM














When you accidentally forget to switch the oil flow rate down after priming

We typically prime all 3 pumps at 30,000 ul/hr, and then change the flow rates back down to what they should be for that particular chip.  It’s crucial to prime (push all the air out of the tubing) using the pump and not by hand, to ensure that the pump is truly flush with the plunger and ready to start the moment you press “Run”.  If you ever forget to adjust the oil back down to the 13,000-15,000 ul/hr range, here’s what the triangle might look like:

Screen Shot 2016-05-09 at 5.56.29 PM


















Example of uniform droplets

Note that our current devices (homemade) produce droplets that are slightly smaller than the company devices.  To the right, you see droplets that are still sitting on top of each other.  To the left, you see droplets that have dispersed into a single layer.  Droplets will typically disperse on their own to facilitate counting if you let them sit for a few minutes, but it is good to check for quality right away because droplets will begin to merge in the C-chip, particularly those on the far edge.  If you see larger droplets distributed throughout (versus only being at the very far edge), that’s an indication of poor droplet quality.

Uniform droplets





















Here’s a different experiment with near-perfect droplet quality, but this was taken after some droplets had begun merging together.  Be gentle when you’re pipetting the droplets into the C-chip, otherwise some droplets might burst immediately. 

Merging droplets






























[coming soon]




What should my libraries look like and how do I get rid of primer dimer?

The yield of your libraries will depend on your specific sample and how many cycles you choose for PCR.  Sometimes both types of libraries will vary a bit with regard to how wide or skinny they are.  However, both traces should be relatively smooth, as shown below.


A typical cDNA trace for a human-mouse run

Usually the amount of primer dimer (a peak that shows up just to the right of the lower marker) will be relatively low compared to your actual library, in which case don’t worry about it and proceed to tagmentation.  If the primer dimer peak is nearly as big as your sample, try doing a second purification with .6X AMPure beads.

cDNA trace










A typical cDNA trace for a primary tissue run

These traces have a few spikes, but note that the entire trace is not a series of spikes – rather we only see a few of them.

primary tissue cdna

primary trace









A typical post-tagmentation trace

To ensure that you’re loading a high quality library onto the sequencer, double purify your libraries by first purifying once with .6X AMPure beads, followed by a 1X purification.

Tag trace










– Melissa Goldman