Milli (Vanilli) Fluidics - Part 2

4% agarose in PBS (200 g/L salinity, pH 7.2)

The idea has been refined:

• Make two adjacent channels with a weir in between
• Cells get caught in the weir as flow moves in from one channel, over the weir, and out the other channel
• Same flow path is used for adding stimulant

Making grooves
Attempt 1
Used two fine gauge wires covered in tape.  Found some wire in the common tool area, from the looks of it, it was 30awg.  The wires provided the depth for the channels while the tape smoothed out the profile and made the weir in between the wires/channels.

The wires were positioned so that neither channel extended completely across the pad region, but with a small (~5-10mm) of overlapping region for the weir.

Wire+tape 'mold' was assembled on a regular office grade transparency sheet to make release of the mold from the agarose pad easier.

Result
Channels made this way were really really deep.  The pad patterned was also too small.  so channels went all the way across.  Didn't test loading with cells.  Regular Scotch tape (matte finish) produced a lot of observable roughness in the resultant patterned pad.  Prefer it to be smooth.

Attempt 2
Instead of wire, used strips of tape, covered in tape.  Much easier to position.  Used clear packing tape as the 'smoothing' layer.

Result
Grooves pattern probably caused by scratches from tweezers used to smooth out air bubbles.  Cells get caught in these grooves.  Cells dropped onto the pad (5uL) before coverslip.  Trapping in the weir doesn't seem robust.

Attempt 3 
Discontinuous single channel patterned using wires heavily smoothed with tape.

Result
Attempted cell loading via channel.  Appears that there is enough non-uniformity (non-flatness) in the pad that promotes bulk capillary action across the entire pad.

Attempt 4
Fine grit sand paper roughened packing tape.  Sandpaper drawn over surface once with moderate pressure, to create deep parallel grooves (will result in channel walls on pad).

Result
The pattern was evident when the pad was dry, but once wetted, it was not observable.  Only 2uL of cells spotted (small pad).  Cells in center popped - probably from over pressure of coverslip.

Todo: Attempt 5
PDMS 'mold box'.  Put a small transparency film roughened with sandpaper on a flat surface.  Outline the film with tape, approx 2-5mm from the film edge.  Cast PDMS over the top of this.

The PDMS will be come a small molding box to create a pad the size of the transparancy film with grooves from the sandpaper roughened area.  Drop this box over a spot of molten agarose on a glass slide.  Cool the assembly and peel away (hopefully with the agarose portion still stuck to the glass slide).

Milli (Vanilli) Fluidics

I'm working on some experiments with algae (while we rush to publication - lest we get scooped - ugh!) that require quickly isolating a few and stimulating them with a grab bag of chemicals.  I have microfluidic devices specifically designed for other purposes, but shoe horning them into this experiment will probably be a project in and of itself.

I've made "poorman's" microfluidics by cutting pieces of scotch tape on glass slides and casting PDMS against it. However, the cells I'm working with are small and motile enough to make anything created this way totally inadequate.  Besides, I'm a bit worried that the random organics in PDMS (and Scotch tape adhesive) can throw off my cellular response.

So here's what I want to test - an agar/agarose pad with a groove patterned it in connected to a Laplace pressure pump.  More specifics:

• I can make a dozen of these pads (at least ones without a groove) in about an hour
• To make the groove I need to pattern the slide I use to 'flatten' the pad
• The Laplace pump will be made with a coverslip with a tiny hole in it that:
• I place a drop of stimulant over
• will be far away from where I image cells
• needs to be aligned to the groove in the pad

Will this work?  Off to the drawing board!