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.
Sunday, February 23, 2014
Trustee Presentation
On Friday (February 21st), I was again unable to attend my internship because it was the day of the Emma Talks! Emma talks offered a very unique opportunity to hear from six of the most impactful women in the world speaking on topics from the environment, to women in the workplace, to the importance of girls' education.
After the Emma Talks, I had the opportunity to present my own work from my internship to the Board of Trustees! It was a great opportunity to share what we're working on.
I look forward to getting back into the lab this week and starting my blocking buffer experiment!
After the Emma Talks, I had the opportunity to present my own work from my internship to the Board of Trustees! It was a great opportunity to share what we're working on.
I look forward to getting back into the lab this week and starting my blocking buffer experiment!
Sunday, February 16, 2014
Snow Day!
On Friday (February 14th), I was unable to attend my internship because we had a snow day! Instead of any scientific updates, here's a beautiful picture of Emma in snow!
Saturday, February 8, 2014
Testing Dye Binding
Yesterday (February 7th), I continued work on my new project. I ran a preliminary test to confirm the way in which dye binds to slides with PBS and Casein. From this experiment, we expected that the slide incubated with casein would be stained, while the slide incubated with PBS would not. To test this, I incubated one slide with 3 mL of 10X Casein and one slide with 3 mL of PBS for an hour on the shaker. Once the incubation was complete, I drained the solutions into the biologic waste container and washed both of the slides with PBS three times for 10 minutes each time. I then added 3 mL of LabSafe GEL Blue stain onto each slide for 20 minutes.
Once this process was complete, we found a surprising result. The PBS slide, as expected, had not bound the dye. However, the Casein slide did not bind the dye either. Instead, the protein had bound the dye, and the protein was no longer bound to the slide. In the image below, the Casein slide (left) and PBS slide (right) are both pictured with the dye solution still on the slides.
After seeing this result, we decided that either a small amount of Casein was stuck under the slide and came up to bind the dye, preventing us from seeing the protein on the surface of the slide, or the dye eluted the Casein from the slide. In an attempt to see if there was still protein bound to the surface of the slide, I poured the dye off of the slide, washed it with PBS, sprayed it with water, and added 3 mL of dye back onto the slide in a different container. After 10 minutes, the slide showed no change in color, leading us to believe that there was no protein bound to the slide. We will test another Casein slide to see if we get the same result. If the dye is found to elute the Casein from the slide, we will need to reconsider the procedure for my project.
While waiting for the slides in this experiment to incubate, I labeled the trays for my project and prepared my slides by washing them in ethanol.
I look forward to finding out if the dye does indeed elute the Casein from the slides.
Once this process was complete, we found a surprising result. The PBS slide, as expected, had not bound the dye. However, the Casein slide did not bind the dye either. Instead, the protein had bound the dye, and the protein was no longer bound to the slide. In the image below, the Casein slide (left) and PBS slide (right) are both pictured with the dye solution still on the slides.
After seeing this result, we decided that either a small amount of Casein was stuck under the slide and came up to bind the dye, preventing us from seeing the protein on the surface of the slide, or the dye eluted the Casein from the slide. In an attempt to see if there was still protein bound to the surface of the slide, I poured the dye off of the slide, washed it with PBS, sprayed it with water, and added 3 mL of dye back onto the slide in a different container. After 10 minutes, the slide showed no change in color, leading us to believe that there was no protein bound to the slide. We will test another Casein slide to see if we get the same result. If the dye is found to elute the Casein from the slide, we will need to reconsider the procedure for my project.
While waiting for the slides in this experiment to incubate, I labeled the trays for my project and prepared my slides by washing them in ethanol.
I look forward to finding out if the dye does indeed elute the Casein from the slides.
Saturday, February 1, 2014
First Step: Blocking Buffer Solutions
On Friday (January 31st), I worked on the first steps of my new blocking buffer project! Because I did not specifically explain the purpose of a blocking buffer in my previous posts, blocking buffers are used to prevent nonspecific binding, reduce background signal, and stabilize proteins for better interactions. These effects are demonstrated in the following image.
Today I took my first step in my project by making my blocking buffer solutions! We decided to make 5 mL of each BSA solution (5%, 2.5%, 1%, 0.2%) and 10 mL of each Casein solution (1x and 1/2x). First, I had to do the calculations to determine the amounts of the buffers and the amounts of PBS that I would need for the dilutions. First, I would make the highest concentration solution and dilute from that. Below are my calculations.
Once the calculations were confirmed, I completed the dilutions. I have now officially started my own project! I can't wait to carry out this experiment!
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| Image from http://www.immunochemistry.com/products/elisa-solutions-1/blocking-buffers.html |
Today I took my first step in my project by making my blocking buffer solutions! We decided to make 5 mL of each BSA solution (5%, 2.5%, 1%, 0.2%) and 10 mL of each Casein solution (1x and 1/2x). First, I had to do the calculations to determine the amounts of the buffers and the amounts of PBS that I would need for the dilutions. First, I would make the highest concentration solution and dilute from that. Below are my calculations.
Once the calculations were confirmed, I completed the dilutions. I have now officially started my own project! I can't wait to carry out this experiment!
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