- Put coverslip with adherent cells in the sample chamber and wash with PBS. This involves gently spraying the PBS on the coverslip with a pippette, and gently rocking the sample chamber back and forth. This should only require about 0.5 mL of PBS.
- Aspirate PBS and fill chamber with 1 mL of 4% formaldehyde. Let sit for 15 minutes at room temperature (in cell prep box, covered with a chem wipe).
- There are variances in both time and temperature (some incubate rather than using room temperature), so this is something that could be varied if desired.
- Aspirate formaldehyde and wash twice with PBS. Aspirate PBS.
- For measurements, fill with HBSS at 37C as is typical with live cell measurements.
Raman Spectra of 1um beads taken at 20s exposure times. Spectra were taken at different axial positions of trapped bead(s?) and a group of beads that were adhered to the cover slip. These positions ranged from about 5 (microns?) above and below the cover slip (full range is shown for individual beads while select points are shown for the ensemble). The “glass edge” on the right is dominant in all of the adhered ensemble spectra while it disappears as the focus moved farther into the sample for the trapped bead.
Some sources of uncertainty are: 1)finding a place where a bead could be trapped and wouldn’t get knocked out by other beads coming into the trap or leave on its own (I had to start 5 microns above the coverslip to get a stable trap). 2)I used the back reflection of the glass on the microscope’s ccd as a reference point. When I was finishing measurements, I noticed the position of the back reflection was not where I originally set my “0” point (This is not the first time this has been noticed, but I wanted to make a note of it here) 3)I’m pretty sure that 1 increment on the fine tuning knob (which is what I was using to move the objective) corresponds to 1 micron, but I haven’t been able to 100% verify this. 4)Looking at published spectra, the pixel to wavenumber calibration may be slightly off (peaks are shifted to the right a bit). 5)This spectra was taken after looking for and reducing sources of power loss in the Raman path. Part of the process involved adjusting a filter that cleans up the laser light and that may explain the floor around 200 cm^-1
Raman spectra of a plastic block taken at different exposure times ranging from 1 second to 2 minutes (range in images shown is 1 to 30 seconds). Increasing exposure time reduced noise but also increased background. Spectra was collected with the oil objective and the block in direct contact with the objective, so changing depth was not an option. I also looked at how the spectra looked using the full CCD (fvb) and a subset corresponding to an individual fiber from the bundle (mt).
These are Raman spectra of ethanol taken using the IRAM system. Images were taken at multiples of 5 microns into the sample, starting from the coverslip/ethanol interface to 25 microns into the sample. As the focus goes deeper in to the sample, the effects of the coverslip (ex broad peak at the high end) become less prominent.
Exposure time was 10 seconds. Spectra was taken from a section of the image that was 10 pixels tall, corresponding to light going into a single fiber of the fiber bundle.
For today’s group meeting, only the IRAM members (Xing, Robert, and Ashley) were in attendance. We primarily discussed the big picture scope of our project and the upcoming Photonics West presentations. Continue reading Group Meeting 10/23/15
To our knowledge, the throughput of our system versus angles has not previously been accounted for. To determine it, we placed the teflon cap in the sample chamber, submerged it in water and put a square coverslip on top, just as we would for a bead sample. The scattergram for the cap is shown below.
- If pellicule beamsplitter is out, put it in between the CCD and the last lens of the 4f system. The engraved side should be facing away from the CCD. It should be at as much of a 45 degree angle as possible, given the space constraints. Make sure the beamsplitter is not clipping the sample beam.
- Direct the reference beam towards the CCD using reference mirrors 1 and 2.
- With all ND filters out, adjust reference mirror 1 so that the reference beam transmitted through the pellicule and the sample beam reflected off of it hit the same spot in the near field (close to the pellicule).
- Adjust reference mirror 2 so that the two spots overlap in the far field (farther away from the pellicule).
- When the spots are very close to overlapping, it may be beneficial to stop down both beams so that the spot sizes are small. Just make sure the iris’s used are centered!
- Put the ND filters back in, and make final adjustments to mirror 1 so that the two beams overlap perfectly.
- If this part of the alignment is correct, you should have circular fringes with good contrast.
- Put a sample of fixed beads into the system (larger beads make alignment easier).
- Place a bead on the edge of the elastic excitation beam, diagonal from the center. Adjust BS1 and reference mirror 1 so that the tilt fringes are nulled in the interferogram.
- Move the bead to the center of the excitation beam. Adjust the BS and mirror until the frequency of the tilt fringes approximately doubles in both x and y directions.
- Put in two pinholes along the reference beam path to make future alignments easier.
Raman alignment (all in epi-illumination mode):
1) With the air objective and no condenser lens, and a glass coverslip in the sample plane, use the 1st Raman mirror to make the two reflections from the coverslip overlap on the hitachi CCD.
2) With no objective or condenser lens, and a mirror facing downwards in the sample plane, use the 2nd Raman mirror to make the retro-reflection exiting the microscope hit the same spot as the illumination beam. (This can be done by using the IR viewer and a piece of lens paper to see both beams.)
3) Repeat steps 1-2 until they converge.
4) Put in the oil objective and a glass coverslip in the sample plane. Adjust the 2nd Raman mirror so that as the objective goes in and out of focus, the beam of the hitachi CCD has equal power in each quadrant of the spot.
5) Put in two pinholes after the periscope. These pinholes designate the path of a vertical beam.
Elastic alignment (in trans-illumination mode):
6) Use the two Elastic mirrors to direct the beam through the pinholes before the periscope, using the 1st mirror to align to the first pinhole and the 2nd mirror to align the second pinhole.
7) To fine-tune the beam so that it is going through the center of the objective with no condenser lens: Adjust the 2nd Elastic mirror so that the beam is hitting the center of the objective (will need IR viewer to see this). Then adjust the 1st Elastic mirror so that the beam is still going through the pinholes before the periscope. Repeat until the two converge.
8) Put in the condenser lens, and adjust its focus and position.
1) In epi-mode: Adjust 1st and 2nd Raman mirrors so that the beam is going through the 1st and 2nd pinholes, respectively, in the Raman excitation path.
2) In epi-mode: Without the condenser or objective, adjust the periscope mirrors so that the beam is going through the two pinholes after the periscope.
3) In trans-mode: Adjust the 1st and 2nd Elastic mirrors so that the beam is going through the two pinholes before the periscope.
4) Put in the objective and condenser. Adjust the focus and position of the condensor.
These days work are mainly about comparing the performance of normal fitting and direct fitting.
A review for whole process. We have a parent Gauss or Normal distribution(parameter μ and σ). In math that should be continuous. Then we sample the distribution got a set of discrete data. Then we use the discrete population to map to a set of scattergram that generate by Mie scatter thoery in different particle diameter. After that we use cross-section to weight the data. Now we get the raw simulation data we use in normal fitting.
In normal fitting we use log-normal population model + cross-section weighted model+Mie theory to fit to the scattergram. We want to report best estimate of I vs d.
For direct fitting, the raw simulation data generate is similar to the former one. We have a parent Gauss or Normal distribution(parameter μ and σ), then we sample the distribution get a set of discrete data. We weighted these data though cross-weighting model. Now we get the raw simulation data.
In direct fitting, we use a log-normal or gaussian population model to fit raw data directly(without Mie thoery, that is why we call it direct fit).
The purpose of normal fitting is we want to analyze cell’s angular scattergram we taking from our experiment devices. We would like to report best estimate of I vs d.
The purpose of comparing these two fitting process is we want to know several things: how well our normal fit compared to the direct fit; when the samples are very sparse, the Mie thoery model’s contribution could be neglected, and so on.
Followings are several simulation results for comparing direct fitting and normal fitting. Continue reading Compare normal fitting with direct fitting
Thawing Frozen Cells
- Cells arrive deep-frozen.
- Add cells to 10 mL of growth media with 20% FBS.
- Incubate overnight or until confluent.
- Passage to 2 dishes with 10 mL of growth media with 10% FBS
- Continue to passage about twice a week.
- Aspirate (remove) media.
- Add 1 mL of trypsin, coating the entire bottom of the dish.
- Swirl for 20 seconds, then aspirate.
- Add 1 mL trypsin.
- Incubate for 4 minutes.
- Use the pipette to “pressure wash” cells off of one dish with trypsin.
- Add the trypsin with the lifted cells to the second dish.
- Use pipette to wash cells off of the second dish with trypsin.
- Add (x) μL of trypsin with lifted cells to new dishes with 10 mL of growth media.
- (x) should be somewhere between 75 and 150, depending on how quickly you want the cells to reach confluence