Innocent Until Proven Guilty: Evaluating Pore Patency Across 0.5 μm Microporous Membranes for use in μSiM-CA Bacterial Assays

When we first began developing a standardized protocol for μSiM-CA bacterial assays (described here), bad assay performance led some to believe our issues were due to availability of pores within the membranes inside of the μSiMs. At the same time, I developed a visualization protocol to view bacterial transmission across the membrane (described here). While imaging membranes in DIC, I would occasionally see pores which looked to be filled with air bubbles. Seeing as the presence of bubbles within pores may decrease pore participation once flow is applied to the membrane and also facing continued pressure to prove the membranes innocence, we decided to check for pore participation using confocal imaging and fluorescent beads. The results from this approach not only helped to produce the pull down modification that we are currently testing in μSiM-CA bacterial assays (described here), but it also cleared up any doubt that pore availability within our membranes may be the source of our troubles with μSiM-CA bacterial assays.

Suspect Number One: Bubbles

Suspicious of bubbles within pores of the membranes, we sought to get an idea of how widespread this might be across the membrane surface. Our fear was bubbles within pores could block fluid flow, effectively knocking them out of participation in any application utilizing flow or transmission across the membrane. We believed that bubbles probably formed during the initial wetting of the μSiM-CA, so we also sought to gauge three different wetting techniques and their ability to limit the creation of bubbles within the membrane. In all of our tests we wet the bottom channel the same way in all devices, described in the following:

‘Pipette ~15 µL of PBS by inserting the pipette tip into one of the two open ports of the µSiM-CA and depressing the plunger of the pipette; the PBS should flow from this port through the bottom channel and out the opposite, open port. Remove the pipette tip before releasing the plunger of the pipette to avoid sucking injected fluid back out of the bottom channel of the device’

The three wetting techniques for the well were fairly similar, each slightly modifying the previous approach. The step in the protocol (linked above) that we were modifying is the following:

‘Fill the well of the µSiM-CA by pipetting 100 µL of fresh PBS into it; care is taken to not create air bubbles and/or remove them by by withdrawing injected media and injecting it again until no air bubbles are visible.’

The first wetting technique involved wetting the well of the μSiM-CA from the side of the membrane (near any of the acrylic walls of the well). The second only changes the location with which wetting is done (from the center of the well, which is also the center of the chip). The last technique is the same as the second, but in addition a 1 minute plasma wand treatment (at max power) is done on the membrane before wetting. Hypothetically, the wettability of the surface increases as we go from the first technique to the last. If this is true and there is a correlation between bubble formation and surface wettability, we should see the most bubbles in the first technique and the least bubbles in the third technique. While we did find bubbles after wetting membranes with all three techniques, shown in Figure 1 below, no technique clearly showed more bubbles than the others.

It was hard to find any bubbles at all within any of the membranes. Because of this it was also challenging to definitively say whether or not one technique was better than the other. Therefore, we decided to take a different approach and use fluorescent beads to highlight any areas that may not be participating in fluid flow through the membrane.

Widening the Search: Withdrawing Fluorescent Beads Onto the Membrane to Visualize Blocked Pores

Opposite of our previous approach (searching for signal from a high background), our idea was this approach would search for signal from a low background which would be easier to detect. It also had the added benefit that at the right concentration, we thought any pores that were occluded for any reason (bubbles, debris, under etched) would show up as dark spots on an otherwise lit up membrane. This technique therefore presented as a better gauge of pore patency across the membranes, with fluorescent beads capturing in open pores but not in occluded pores. The number of pores across a microporous membrane is variable, but we estimate that there are about 1E6 per membrane window (after counting by hand). In our three window chip, this means there are roughly 3E6 pores across the entire chip. As a result, we decided that loading the well with 8.75E6 510 nm diameter beads and withdrawing 50 uL from the total well volume of 100 uL, 4.375E6 beads total, would provide every pore with the opportunity to have a bead in it if it were open. As we did before, we tested three different wetting techniques to see if bead capture would be affected. To get beads onto the membrane surface, we used a pipette to withdraw our bead solution onto the membrane. This protocol is as follows:

Protocol

1. Pipette ~15 µL of PBS by inserting the pipette tip into one of the two open ports of the µSiM-CA and depressing the plunger of the pipette; the PBS should flow from this port through the bottom channel and out the opposite, open port. Remove the pipette tip before releasing the plunger of the pipette to avoid sucking injected fluid back out of the bottom channel of the device
2. Fill the well of the µSiM-CA by pipetting 100 µL of fresh PBS into it; care is taken to not create air bubbles and/or remove them by by withdrawing injected media and injecting it again until no air bubbles are visible
3. With the bottom channel and well wet, block one of the open ports using a 3M double-sided tape sticker and stabilize the devices with clamps around its sides
4. In the same manner as above, pipette ~40 µL of PBS into the open port and look to see the well fill with little resistance. If resistance is appreciable, discard the device and prepare a new one
5. Remove all liquid from the well (~140 µL)
6. Next, add 100 µL of the bead solution to the well of each device, withdrawing and injecting the bead solution again if air bubbles are present until they are gone
7. Withdraw 50 µL of fluid from the open port, sucking fluid from the well through the membrane into the bottom channel and out the port
8. Image on the confocal microscope

Our results showed us that our pulldown technique worked, but it also showed us that our bead concentration was too low. The results are shown in Figure 2 below.

Having our bead concentration so low not only made it difficult to discern any difference between the three wetting techniques, but it also made it challenging to determine if there were any occluded pores. As a result, our next step was to repeat the experiment but increase our bead concentration.

Fine-Tuning Our Approach: Increasing Bead Concentration for Better Visualization

Seeing that withdrawing 4.375E6 beads onto the membrane was too few beads, we increased the bead concentration within the well nearly two orders of a magnitude to 1.359E8 beads. This means that with a 50 uL withdrawal we would be pulling 6.795E7 beads down on to the surface of the membrane. Using the same protocol as before, we obtained the results shown in Figure 3. This concentration allowed us to visualize areas of pores which were occluded on the membrane, but the creation of a thick cake of beads on top of the membrane prevented us from gauging any difference between the three wetting techniques past this.

Similar amounts of bare spots were seen across all three wetting techniques, but the total area of the bare spots compared to total area of the rest of the membrane was very low. This seemed to indicate an overwhelming majority of the pores across the membrane were in fact participating in fluid flow and subsequent bead capture. To get a better idea of how the three different wetting techniques may affect the abundance of these bare spots, we slightly decreased our bead concentration for our next attempt. Instead of loading the well with 1.359E8 beads, we loaded the well with 5E7 beads. This means that with a 50 uL withdrawal, we would be pulling 2.5E7 beads down on to the surface of the membrane. These results are shown in Figure 4 below and are very similar to our previous results.

This concentration of beads looked better, forming less of a cake as the previous concentration but still blanketing the surface of the membrane with a generous amount of beads. Once again it was challenging to find pores on the membrane that did not have beads in them, regardless of which wetting technique we used. When I did find areas without beads, they were very small and relatively insignificant compared to the rest of the membrane which was able to capture beads.

Conclusion

These results finally allowed us to conclude that a) an overwhelming majority of the pores (>99%) participate in flow and are not occluded by bubbles, debris, or are significantly under etched and b) no matter which technique is used to wet the membrane, bubbles within the pores do not seem to be a significant issue or contributor to poor assay performance while using the μSiM-CA. This evidence therefore suggests that, with regard to membrane structure and pore availability, poor assay performance is unrelated to the membrane since greater than 99% of the pores are open and participating in fluid flow.