Friday, March 21, 2014
Missed Weeks
Last Friday (March 14th), my mentor was on spring break, so I did not go to RPI. For the next two weeks (March 21st and March 28th), I will be on spring break, so I will not be going to the lab. I look forward to returning on April 4th!
Sunday, March 9, 2014
Imaging Room Temperature Slides
On Friday (March 7th), I took images of the slides from my room temperature experiment from last week! I started by taking white light pictures with my phone. I took one image per blocking time, with each concentration labeled. I also included the control slide in each image, which was blocked with PBS for 2 hours. Here are the resulting images.
The staining was opposite of what we initially expected- the lowest concentrations were stained darkest.
After finishing the white light images, I went down to the flatbed scanner to scan the slides. Even though this dye does not really fluoresce, we scanned them for fluorescence anyway to get images. Each column on the scanner corresponded to a certain time, with the concentration increases up the column.
In the resulting image, the darkest-stained slides turned out as the lightest when they were scanned, so we had to invert the resulting images, so the darkest-stained slides show as the darkest. I will eventually use these scanned images to evaluate the darkness of each slide in pixels to quantify what is otherwise qualitative data. This imaging will be the data to take away from the experiment as a whole, to determine the blocking buffer, time and temperature that will maximize blocking efficiency in other experiments.
JP is on spring break next week, and then I am on spring break for the two following weeks, so I will not be able to return to RPI until April 4th. I can't wait to get started on the next phase of my project, which will be the 37C experiment!
The staining was opposite of what we initially expected- the lowest concentrations were stained darkest.
After finishing the white light images, I went down to the flatbed scanner to scan the slides. Even though this dye does not really fluoresce, we scanned them for fluorescence anyway to get images. Each column on the scanner corresponded to a certain time, with the concentration increases up the column.
In the resulting image, the darkest-stained slides turned out as the lightest when they were scanned, so we had to invert the resulting images, so the darkest-stained slides show as the darkest. I will eventually use these scanned images to evaluate the darkness of each slide in pixels to quantify what is otherwise qualitative data. This imaging will be the data to take away from the experiment as a whole, to determine the blocking buffer, time and temperature that will maximize blocking efficiency in other experiments.
JP is on spring break next week, and then I am on spring break for the two following weeks, so I will not be able to return to RPI until April 4th. I can't wait to get started on the next phase of my project, which will be the 37C experiment!
Sunday, March 2, 2014
Blocking Buffer Room Temperature Phase
On Friday (February 28th), I finally started my project for real! Today I ran the room temperature phase. At room temperature, Casein and BSA were each blocked for times of 30 minutes, 1 hour, and 2 hours- Casein at 1/2x and 1x and BSA at 0.2, 1, 2.5, and 5%. The slides I ran today are indicated in yellow on the schedule below. I also added a control slide that was blocked with PBS for 2 hours.
My protocol was as follows:
1) Add 1mL of the specified protein solution to the specified slide.
2) Block on the shaker for the specified time.
3) Drain well with pipette into waste container.
4) Wash slide with 4 mL PBS for 5 minutes.
5) Drain well with pipette into waste container.
6) Transfer slide to new plate.
7) Stain with 4 mL blue stain for 15 minutes.
8) Drain stain into waste.
9) Wash slide with 4 mL PBS for 5 minutes.
10) Air dry slides for 5 minutes on paper towel.
11) Write blocking solution, blocking time, and temperature on back of slides.
When staining, the blue dye clotted with 5% BSA blocked for 30 minutes, 5%, 2.5%, and 1% BSA blocked for 1 hour, and 1% BSA blocked for 2 hours. In the future, we will work to find the cause of this clotting.
In this image, the 1 and 2 hour blocking time slides are still blocking with their respective protein solutions (step 2). The 30 minutes blocking time slides are being stained with blue dye (step 7). You can see the clotting of the 5% BSA slide on the far right.
I also had a photographer in the lab today, which added to the excitement of the day! I can't wait to continue my experiment in the coming weeks and begin to analyze the results.
My protocol was as follows:
1) Add 1mL of the specified protein solution to the specified slide.
2) Block on the shaker for the specified time.
3) Drain well with pipette into waste container.
4) Wash slide with 4 mL PBS for 5 minutes.
5) Drain well with pipette into waste container.
6) Transfer slide to new plate.
7) Stain with 4 mL blue stain for 15 minutes.
8) Drain stain into waste.
9) Wash slide with 4 mL PBS for 5 minutes.
10) Air dry slides for 5 minutes on paper towel.
11) Write blocking solution, blocking time, and temperature on back of slides.
When staining, the blue dye clotted with 5% BSA blocked for 30 minutes, 5%, 2.5%, and 1% BSA blocked for 1 hour, and 1% BSA blocked for 2 hours. In the future, we will work to find the cause of this clotting.
In this image, the 1 and 2 hour blocking time slides are still blocking with their respective protein solutions (step 2). The 30 minutes blocking time slides are being stained with blue dye (step 7). You can see the clotting of the 5% BSA slide on the far right.
I also had a photographer in the lab today, which added to the excitement of the day! I can't wait to continue my experiment in the coming weeks and begin to analyze the results.
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