The experiment platforms for the microfluidics laboratory consist
of a microfluidic chamber and a video microscope that is controlled
via a computer.
Topologically, our experiment chamber is a long (approximately 30 cm)
slender (250 micrometers wide, 75 micrometers high) channel that runs
between five reservoirs that can be filled with test solutions and cells.
If the two-reservoir end is used as the input, then the solutions
in those reservoirs will flow side-by-side as they enter the central channel.
Such flow is said to be "laminar" because the fluids seem to be in layers.
As the fluid progresses down the channel, the fluids will gradually mix
Such mixing is illustrated in the following photograph.
Alternatively, the three-reservoir end can be used as input,
in which case the flow will be "tri-laminar."
This configuration can be useful for mixing cells contained in the
central reservoir into a test solution contained in the other two reservoirs.
Establishing Fluid Flow
A variety of methods have been used to push fluids through microchambers,
including macroscopic pumps (e.g., syringe pumps, peristaltic pumps, etc.)
and electrically driven flows (e.g., electro-osmosis or electrophoresis).
In this lab, we recommend that you use gravity flow.
With our channel design, a few millimeter difference in fluid level
is sufficient to drive flow rates on the order of micrometers
per second, which are satisfactory for most experiment designs.
Input and output ports connect the microfluidic channels to macroscopic
fluid reservoirs, as shown in the following photograph.
While gravity feed is simple to implement, it can be difficult to start.
If the channel is completely dry, flow should start spontaneously
when the first reservoir is filled.
However, if the channel is filled with a fluid, then air bubbles and
debris can stop the flow.
We recommend the following procedure for filling your chamber with fluid.
- Half-fill all of the reservoirs but one with the desired fluids.
- Snap the remaining reservoir out of the silicone rubber base.
Fully insert a rubber plunger (provided) into the reservoir so that the
rubber end of the plunger is flush with the end of the reservoir.
- Snap the reservoir with plunger back into the silicone rubber base.
Pull on the plunger to gently pull liquid through the channel.
Alternatively, you can attempt to fill stubborn chambers by pushing
fluid through an input reservoir.
Be aware however that excessive positive pressure can break the bond
between the silicone rubber and the underlying glass slide.
Breaking the bond will make the chamber unusable.
Before leaving the lab, please flush your chamber with deionized water
and store the chamber with all reserviors filled with dionized water.
This simplifies starting the flow when the chamber is next used.
The microfluidics chambers can be held in place on the stage of the microscope
The stage has x and y control knobs plus a calibrated focus control knob for z.
The microscope illumination system can be turned on with a toggle switch on
the back of the microscope.
The intensity of the illumination can be adjusted with a control on
the right side of the base of the microscope.
The microscope has 3 objectives: 4X, 10X, and 20X,
which focus an image of the chamber onto a
Measuring Flow Rates
Quantitative interpretation of data from microfluidic
systems often requires knowledge of the flow rates.
Although flow rates can in principle be determined by measuring the time
it takes for some known volume of fluid to flow through system,
it is often more convenient to measure flow rates by tracking
microscopic particles that are suspended in the fluid.
Microscopic polystyrene beads with diameters on the order of 1 micrometer
are available for this purpose.
Because of fluid viscosity, the flow profile across the channel is
approximately parabolic, with faster flow near the center than near the
For that reason, we use beads that are neutrally bouyant in water
(by virtue of a tiny air bubble in each bead!).
If used in solutions with high concentrations of solutes, these
beads can float to the top of the channel (where the fluid moves
much less quickly).
A computer running Linux is attached to the video camera via
You will use a program called
to collect and analyze data from this camera.
What follows are some instructions to get you started.
This program was designed for this lab, and has many important features
that are not described below.
To take full advantage of everything this program offers, we recommend
you read further information on camscope.
Here are a few of the things you an do with camscope:
- Capture images from the camera
- Capture sequences of images (movies)
- Play back previously recorded images and movies
- Measure distances
- Measure velocities
- Measure brightnesses
- Measure cell sizes
To start, log in to the computer using the username camscope and
the password camscope.
Once you log in, there should be a console (a window with a command prompt)
If not, click on the icon that looks like a monitor to open one.
Since multiple groups will be using each computer, you will need to use
directories to keep your data separate.
You can name this directory anything you want, as long as the name is
To make a directory and enter it, type commands like the ones below into
> mkdir I_Love_Camscope
> cd I_Love_Camscope
Once you are in your own directory, type
to activate the main program.
When camscope starts, it will look like the image below.
This is the entire interface: no menus, no pop-up windows, no miraculously
appearing or disappearing buttons, no monkeys to punch.
There are a few controls that don't have visible buttons, but not many.
These are documented at the link provided above.
The main part of the screen is taken up by the image.
This image is either a live feed from the camera, or one of the images you
The text above the image tells you the current image number (every time you
record a new image, the image number goes up by one) and the time at which
the image was taken.
Above that are the data set controls and the camera settings (more about
Below and to the right of the image are two plots.
When the cursor is over the image, you will see a horizontal line and a
The plot below the image displays the RGB (Red, Green, Blue) values for the
pixels along the horizontal line.
The plot to the right of the image displays the RGB values along the
Text between the two plots tells you the current location of the cursor and
the RGB values for the pixel below the cursor.
The text across the bottom reports distance and velocity measurements.
Buttons on the right provide the following functions (top to bottom, with
keyboard shortcuts in parentheses. For all keyboard shortcuts, hold down
the Ctrl key and press the other key listed):
- EXIT (ctrl-q): quit camscope and return to Linux
- PLAY (ctrl-p): display live images from video camera, or loop
through previously recorded images if the program is in Analyze mode.
- PAUSE (ctrl-z): stop displaying new images and hold last one
- RECORD (ctrl-r): start recording images. Records ten images (by
default) and then stops.
- SNAPSHOT (ctrl-s): record a singe image.
- STOP (ctrl-b): stop recording.
- Analyze (ctrl-a): toggles between showing live video and showing
previously recorded images (Analyze mode).
- BACK (ctrl-left_arrow): display previous recorded image
(only works in Analyze mode).
- NEXT (ctrl-right_arrow): display next recorded image
(only works in Analyze mode).
- Screen Capture (ctrl-d): save image of screen (screendump).
There are a variety of measurements you can make with camscope.
We will describe a few here. You can make any of these measurements in
real time from the live camera feed, or from pre-recorded data in Analyze
- Measuring brightness is easy. Simply put the cursor over a pixel in
the image, and camscope will tell you the brightness of that
pixel, along with plots of the brightness of all of the pixels in the same
row or column. If you take a snapshot (ctrl-s) or record a movie (ctrl-r)
while the cursor is over the image, this brightness information will be
saved into a text file along with the image (in live camera mode only).
Note that you'll need to use the keyboard shortcuts for taking images if
you want to do this. You can also measure the average brightness in a
region of the image by clicking and dragging the right mouse button to draw
a box around that region. This average brightness will be displayed next
to the single-pixel brightness data.
- To measure the distance between two points in an image, click and drag
from the starting point to the end using the left mouse button. The
distance will be displayed (in pixels) at the bottom of the screen.
- You can measure velocity the same way you measure distance. However,
because velocity involves time as well as space you will need to combine
information from two or more images. In real-time this is easy — as
you click and drag, new images are being displayed, so you can (for
example) follow a cell with the mouse in real-time to measure its
velocity. In Analyze mode, you will need to switch images manually. For
example, you can click on a cell in one image, and while holding down the
left mouse button press ctrl-right_arrow to switch to the next image. If
you then drag the mouse to the new location of the cell, camscope
will report the velocity of the cell as well as the distance traveled.
- You can measure the area of an object within camscope using a
rudimentary segmentation algorithm built in to the program. When you click
the middle mouse button on a pixel, camscope selects all of the
similar pixels that are in contact with the first pixel and displays them
in magenta. It also reports the size of the selected area (in pixels)
below the Screen Capture button. The algorithm is not perfect, but
there are ways to adjust the settings to fine-tune its performance.
Using Multiple Data Sets to Organize Your Measurements
While you're in the lab, you're likely to want to perform more than one
experiment. If you just collect a bunch of images without organization and
go back to analyze them later, you'll have trouble remembering which images
correspond to which experiment — and you might make mistakes in your
analysis as a result. However, it can be a pain to remember to write down
the image numbers corresponding to each experiment. This is where data
sets come in handy.
Data sets are a convenient way to organize your data by experiment. In
the upper left corner of the camscope screen you'll see a number
that tells you the current data set, along with + and -
buttons (with keyboard shortcuts ctrl-. (period) and ctrl-, (comma)
respectively). If you increment the data set each time you start a new
experiment or change viewing locations, you will only need to write down
the data set rather than the image numbers. Starting a new data set resets
the current image number to zero. All of camscope's other
features (record, analyze, playback, etc) operate on only the current data
set. You have 1000 potential data sets to choose from, so take advantage
Saving Your Results
As you record images, camscope
will save them to the hard drive.
If you quit and restart the program, it will continue recording where it
left off (i.e., it won't overwrite the data you've already taken).
When you are ready to leave the lab, you should save your
data. We cannot guarantee that your data will be preserved on the lab computer,
so copy everything before you leave. The easiest way to save your data is to
record it on a CD. Put a blank CD into the CD burner and type
at the linux prompt. This
command will bring up a graphical interface that lets you save your data to CD.
If your directory is larger than the size of one CD (about
700 MB), you may find it helpful to split your data into multiple
subdirectories and copy each onto a separate CD. Your TA can help you do
IMPORTANT: along with each image is a corresponding text file. If
you delete these text files, or do not save them along with your images,
camscope may not be able to load your saved images later. Be sure
to keep these text files!
is intended to provide sufficient information for you to monitor how
your experiment is proceeding while you are in the lab. This is important so
that you can discover problems and fix the problems before taking your final
data. It is also important so that you have a good idea that your experiment
was successful or not before you leave the lab. If you discover problems with
your data, it is generally easier to take new data now than it is to come back
at a later time and start again.
Results from camscope can also be directly incorporated into your
However, quantitative statistics to test specific hypotheses generally
require additional analyses, based on images and brightness profiles
saved by camscope.
You can generate statistics from the info files by importing those
numbers into text processing, spreadsheet, and/or plotting programs.
You can generate statistics directly from the image files using any
programming environment that supports the .bmp format (e.g., Matlab).
Laboratory Equipment and Supplies
In addition to the microfluidic platforms, the lab also has
- a scale to weigh solutes
- micropipeters to measure small quantities of fluid
- pipets to measure larger quantities of fluid
- disposable pipets to transfer fluids to and from reservoirs
- small plastic bottles with lids to hold and store test solutions
- chemicals: NaCl, KCl, CaCl2, dextrose, sucrose
- food coloring to visualize flows
- deionized water
- and NOTHING else (well actually, there is also air to breathe
— but we don't supply this, so if you use it all up it's your
responsibility to get more).
You are welcome to use additional supplies and equipment to implement
are responsible for obtaining any supplies or equipment
not explicitly listed above.
The staff is only responsible for supplying the equipment described above.
In this experiment, you may use dyes and other chemicals that could stain
or irritate your skin. Wash your hands thoroughly immediately after exposure to
any chemicals. Clean up minor chemical spills immediately. Report major spills
to the staff. Rubber gloves will be available.
No foods or beverages will be allowed in the laboratory.