µSiM in situ Small Molecule Permeability (Confocal Microscope)

Some notes for researchers prior to beginning experimentation: The goal of this experiment is monitor in real time the amount of a fluorescent small molecule that passes a cell layer and enters the bottom channel of the µSiM. This enables us to determine permeability of the cells. See 10 kDa Small Molecule Permeability Optimization for background information. This method should be compatible with both trench-up and trench-down devices, however, only trench-down devices have been used thus far.

Materials

• Appropriate cell line grown to confluency on µSiM device (see appropriate µSiM Device Cell Culturing Protocol)
• Appropriate Cell Culturing Media
• Appropriate fluorescent small molecule
• Pipettes (P200, P20) and tips
• Amber 1.5 mL Eppendorf tubes (or cover with foil)
• Andor Dragonfly Confocal microscope (or equivalent)
• CytoVU (SiMPore)
• Beakers for tip waste and media waste

Methods

Prepare media and FITC Dextran dilutions

1. Warm media in 37ºC water bath.
2. Make fluorescent dye source solution. Protect from light.
   1. Note: Concentrations will need to be optimized on different scopes for proper dynamic ranges and to avoid photobleaching during experimentation. The molecules we have optimized for the Andor Dragonfly Confocal microscope are as follows:
      1. 10 kDa FITC-Dextran: 1 mg/ml
      2. 10 kDa Dextran-AF488: 200 µg/ml
      3. Lucifer Yellow: 150 µg/ml

Set up device and take brightfield images of monolayers

1. Remove old media from device and wash one to two times with media, top and bottom channels.
2. Take brightfield image of cell monolayer.
   1. For optimal phase imaging, add media to form an excess layer on the top of the device (about 115 µL). Carefully drop a coverslip atop the media to form a flat interface.

Prepare confocal microscope for image acquisition

1. Use 10X objective.
2. Optimize channel settings for fluorescent dye.
   1. Dye should be in linear range, no photobleaching should occur over the course of the experiment, dynamic range should be maximized. For instructions on how to optimize a new fluorescent dye, see this page.
   2. The settings we have optimized for the Andor Dragonfly Confocal microscope are as follows:
      1. FITC: 488 nm excitation for light source, 525/50BP nm emission, 0.5 s exposure, 5% laser power
      2. AF488: 488 nm excitation for light source, 525/50BP nm emission, 1 s exposure, 5% laser power
      3. LY: 405 nm excitation for light source, 525/50BP nm emission, 1 s exposure, 35% laser power
3. Set up protocol.
   1. “Time Series” -> “Time Repeat”: repeat all steps every 1 min, 11 times (Take one picture at time 0, and a picture every minute for 10 minutes of diffusion).

Take “in focus” image

1. Wipe the bottom of the device with ethanol and place on microscope in CytoVU.
2. Select “Live”.
3. Find device and focus, placing view in middle of edge, away from corners. Center window. Try to get the membrane is straight as possible in the field of view.
4. Focus on cells and then on membrane edge using microscope knob. The membrane edge should be as crisp as possible.
5. If possible, draw box around membrane edges (transparent region) to enable quick and consistent repositioning of device.
6. Name file (ex: Treatment group_Dev#_if (for “in focus”).
7. Take image for reference to the membrane position.

Add fluorescent dye and take timelapse images

1. Name file (ex: Treatment group_Dev#_test).
2. Remove media from well and quickly reposition on microscope. Get edge crisp again. Alternatively, you can remove the dye prior to taking the “in focus” image. This leaves the cells dry for a longer period of time, and is only recommended if you can do these steps very quickly (i.e. are experienced in the protocol)
   1. This can either be done on the microscope or the device can be removed for pipetting and replaced on the scope.
   2. Realign membrane with rectangle drawn earlier. Move quickly as cells are dye.
3. Drop objective 100 µm.
4. Get ready to start timelapse by placing mouse over “Acquire” or equivalent button to start taking images.
5. Add 100 µl of dye to top well and immediately start timelapse.
   1. After first image is acquired, make sure everything looks normal.
   2. Let it run 10 min. You will see dye entering the bottom channel over time.
      

Take source fluorescent concentration image

1. Name file (ex: Treatment group_Dev#_sc (for “source condition”)).
2. Pipet 20 µL of source dye solution (solution from the well) into channel of device. Repeat to ensure full replacement of media with source solution.
   1. It is easiest to remove device from scope for pipetting. Then reposition on scope, first focusing on edge, then dropping 100 µm below membrane. Always make sure to check for bubbles under the membrane at this step.
      1. Realign membrane with rectangle drawn earlier.
   2. The dye should uniformly flood the channel. For trench down devices, you are focused within the trench, and will only see dye in that region, not in the silicone support region.

Data Analysis and Determination Endothelial Permeability

1. This method may be adjusted, but currently we are using a Mathematica Code that fits data to a “Constant Flux” equation to process images – Contact McGrath lab for details and access to the code.
2. Please reference these posts for background information and theory behind our code:
   1. Theoretical Underpinnings of Small Molecule Permeability Measurements in the µSiM (Part 1: Approach)
   2. Theoretical Underpinnings of Small Molecule Permeability Measurements in the µSiM (Part 2: Experimental Validation)
   3. Theoretical Underpinnings of Small Molecule Permeability Measurements in the µSiM (Part 3: Application to Cell Barriers)*
   4. in situ Permeability Analytical Solution Summary; Unhindered vs Hindered D