Tuesday, May 13, 2014
Reflection
I can't believe this is my final blog post! My experience working in the lab at RPI with Dr. Karande and JP Trasatti for the last two years has been one of the most influential experiences of my life. I have had the opportunity to work on parts of the broader blood-brain barrier project as well as on my own focused blocking buffer solution project. Throughout the last two years, I have been able to learn a wide range of different skills and processes and learn a lot about biochemistry. Having my own project within the lab was especially fun because I got to plan the details of my own project and carry out all the tasks from preparation to procedure to analysis of the results. Looking back, I can't imagine my high school experience without research. The skills I have learned in the lab have crossed over into my academic classes, and doing research has allowed me to expand my learning outside the classroom. The people I have worked with in the lab have been invaluable mentors for me as I have prepared to transition from high school to college and considered what I would like to do in the future. Although work in the lab is very different from regular schoolwork, original research has quickly become something that is of utmost importance to me as I continue my academic career in college and beyond. In fact, my research at RPI has already helped me get a research grant at Washington University in Saint Louis, where I will start in the fall. I'd like to give heartfelt thanks to everyone involved in making my two years in the lab such an incredible experience.
Tuesday, April 29, 2014
Blocking Buffer Results!
Last Friday (April 25th), I finally analyzed my results from the room temperature and 37C runs of my blocking buffer experiment!! First, I took pictures of all of the slides in groups of same blocking temperature and same blocking time. To quantify the results of this experiment, the slides were scanned for fluorescence with a 488nm laser. This imaging resulted in the images below, in which the shading of the slides are actually inverted, i.e. the darkest slide in this image is actually the lightest slide.
The resulting images were then analyzed for mean intensity of each slide using a program called ImageJ. This data was then adjusted to account for the inversion of the images and the background noise of the control slide. The following images show graphical representations of the mean intensity/slide area, where intensity indicates lack of dye, or the amount of protein bound to the slide.
From this data, the optimal blocking conditions for BSA are 2.% for 1 hour at room temperature. There is a general trend for BSA that the amount of protein bound for a given temperature and time period is maximized at 2.5% BSA and decreases for 5% BSA. The only exception to that trend is this trial was the 37C 1 hour data, which may be due to splotching on the slide affecting the mean intensity measurement. The optimal blocking conditions for casein are 1x for 2 hours at room temperature. The amount of casein bound increases with concentration at all time periods at room temperature. Casein does not bind to the slide at 37C.
I can't wait to present my work to the Emma community at the student achievement assembly on April 30th!
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| Scanner image for room temperature slides |
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| Scanner image for 37C Slides |
The resulting images were then analyzed for mean intensity of each slide using a program called ImageJ. This data was then adjusted to account for the inversion of the images and the background noise of the control slide. The following images show graphical representations of the mean intensity/slide area, where intensity indicates lack of dye, or the amount of protein bound to the slide.
From this data, the optimal blocking conditions for BSA are 2.% for 1 hour at room temperature. There is a general trend for BSA that the amount of protein bound for a given temperature and time period is maximized at 2.5% BSA and decreases for 5% BSA. The only exception to that trend is this trial was the 37C 1 hour data, which may be due to splotching on the slide affecting the mean intensity measurement. The optimal blocking conditions for casein are 1x for 2 hours at room temperature. The amount of casein bound increases with concentration at all time periods at room temperature. Casein does not bind to the slide at 37C.
I can't wait to present my work to the Emma community at the student achievement assembly on April 30th!
Saturday, April 12, 2014
Blocking Buffer 37C Phase
On Friday (April 11th), I ran the second phase of my blocking buffer experiment! This consisted of blocking each solution at an incubation temperature of 37˚C. Due to a lack of cut slides, I slightly modified the solutions for this run. For this phase, I used 1/2x and 1x casein at 30 minute and 1 hour blocking times and 0.2%, 2.5%, and 5% BSA at 30 minute and 1 hour blocking times. I gathered all of my materials and went to the 37˚C room down the hall.
For this phase, I used the same protocol from the room temperature phase I ran on February 28th, as seen below.
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.
I will be away on college trips next week, so I will not be able to go to the lab Friday April 28th. I look forward to imaging the slides from this phase when I get back to the lab on April 25th!
For this phase, I used the same protocol from the room temperature phase I ran on February 28th, as seen below.
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.
I will be away on college trips next week, so I will not be able to go to the lab Friday April 28th. I look forward to imaging the slides from this phase when I get back to the lab on April 25th!
Sunday, April 6, 2014
Back to the Lab!
On Friday (April 4th), I finally returned to the lab at RPI after a month of break! Today I took a break from my project to get some experience with MALDI. MALDI is matrix-assisted laser desorption/ionization. This process uses mass/charge ratio of ions to see if a peptide was synthesized correctly. First, I put peptides in solution by adding 100 microliters of acetonitrile-water mixture to each of the six sample. I then added 10 microliters of CHCA and 10 microliters of peptide solution to smaller vials.
We then spotted 1 microliter spots onto a steel MALDI plate. Spots A1 and B1 were CHCA and standards, spots A2 and B2 were sample one, spots A3 and B3 were sample 2, etc.
In the MALDI machine, a laser is used to strike a small area of the sample spot. The sample then ionizes into gases. In the system we are using, differently charged plates serve to shoot the ions down the tube, where there are other differently charged plates at the end. Depending on the size and charge of the ions, the ions go different distances into the field of plates at the end of the tube before they are reflected back towards the reflection detector. Because larger ions go deeper into the end plate area, they take longer to get back to the detector, and the detector can give information about the mass/charge ratio of the ions. This process can be seen in the diagram below.
To shoot the laser, you first choose a part of the sample to shoot because the laser area is much smaller than the area of the spot. The computer program magnifies the sample plate and shows the peptide as white spots within the sample spot. You take multiple samples of each spot to minimize the baseline and increase the signal from the laser.
After finishing the MALDI process, I helped JP check on the brain cell samples and check the CO2 content of the incubator. To check the CO2 level, we used a device that measures the CO2 with a KOH solution. This device attaches to a connector on the incubator with a tube. There is a pump on the tube that you squeeze 18 times to pump a sample of the incubator air into the device. The device is then rotated so the gas combines with the KOH solution. The CO2 dissolves in the KOH solution, increasing the volume of the liquid in the device to a measurable level. We want the concentration to be at 5.0 in the incubator, and we found it to be 5.2, so we recalibrated the incubator to decrease the amount of CO2.
We then spotted 1 microliter spots onto a steel MALDI plate. Spots A1 and B1 were CHCA and standards, spots A2 and B2 were sample one, spots A3 and B3 were sample 2, etc.
In the MALDI machine, a laser is used to strike a small area of the sample spot. The sample then ionizes into gases. In the system we are using, differently charged plates serve to shoot the ions down the tube, where there are other differently charged plates at the end. Depending on the size and charge of the ions, the ions go different distances into the field of plates at the end of the tube before they are reflected back towards the reflection detector. Because larger ions go deeper into the end plate area, they take longer to get back to the detector, and the detector can give information about the mass/charge ratio of the ions. This process can be seen in the diagram below.
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| Image by JP Trasatti, 4/4/2014 |
After finishing the MALDI process, I helped JP check on the brain cell samples and check the CO2 content of the incubator. To check the CO2 level, we used a device that measures the CO2 with a KOH solution. This device attaches to a connector on the incubator with a tube. There is a pump on the tube that you squeeze 18 times to pump a sample of the incubator air into the device. The device is then rotated so the gas combines with the KOH solution. The CO2 dissolves in the KOH solution, increasing the volume of the liquid in the device to a measurable level. We want the concentration to be at 5.0 in the incubator, and we found it to be 5.2, so we recalibrated the incubator to decrease the amount of CO2.
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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