Investigating the Interactome of TMEM106B Monomers vs. Dimers in Frontotemporal Dementia Using Proteomics and Bioinformatics Pipelines
Advisor: Karen Allendoerfer
Internship Location: Stanford University
Onsite Mentor: Stephanie (Steph) Rayner; postdoc; Stanford University
TMEM106B is a key genetic risk factor in frontotemporal dementia (FTD). This gene encodes for the lysosomal transmembrane protein TMEM106B and often forms polymorphic dimers with itself (“homodimers”), with high levels protecting against the disease and lower levels leading to increased risk of FTD. For this project, I am conducting wet lab experiments at the Gitler Lab using a HEK293T (human embryonic kidney) cell model to determine how TMEM106B’s protein interaction partners differ between TMEM106B monomers and dimers. Specifically, I am using transfection to introduce these plasmid constructs into cells and immunoprecipitation to pull out my desired proteins, then sending them to a mass spectrometry facility in Australia and analyzing my results via bioinformatics tools. This research will provide new molecular insights into whether the monomerization of TMEM106B may affect its protein interaction partners and further elucidate mechanisms underpinning FTD.
Project Blog
Week 0: Introduction
Hi, I’m Amanda! Welcome to the first week of blogging my senior project.
This project originated when I brought up the possibility of pursuing a senior project with the lab I have been working with the past few months, the Gitler Lab. The Gitler Lab focuses on investigating the genetic factors contributing to ALS and FTD. Being in the lab has allowed me to develop an interest in ALS/FTD, and this is a field I may continue to dive deeper into in my path toward medicine. Before I get started with my actual project, I wanted to go over some background on the objectives of my project. At the very end of this post, I have also included a glossary for reference.
Amyotrophic lateral sclerosis (ALS) and frontotemporal dementia (FTD) are closely-related, debilitating neurodegenerative diseases. For this project, I will be comparing the protein interaction partners between monomer and dimer formation within the gene TMEM106B, a key genetic risk factor in FTD/ALS. TMEM106B encodes for the lysosome transmembrane protein TMEM106B and often forms polymorphic dimers with itself (“homodimers”), with high levels protecting against the diseases and lower levels leading to increased risk.
In more than 97% of ALS patients and 45% of FTD patients, the protein TDP-43 is found maladaptively depleted from the nucleus and aggregated in the cytoplasm. Its mislocalization leads to a loss-of-function in the nucleus but a gain-of-function in the cytoplasm, where it forms stress granules. Within the nucleus, recent research has shown TDP-43 to also be essential in regulating alternative polyadenylation (APA), an RNA processing pathway that describes the usage of different polyadenylation sites and is found in more than 60% of human genes. Our lab was among the first to discover that APA could lead to RNA sequences with different 3’-UTR lengths. The 3’-UTR is the regulatory region in mRNA immediately after the stop codon and includes most polyA sites (in my gene of interest, TMEM106B, all polyA sites are in this region). As binding to distal polyA sites leads to longer 3’ UTRs compared to proximal polyA sites, APA site usage alters 3’-UTR lengths. When TDP-43 is depleted, TMEM106B transcripts lengthen. This seems to decrease dimer formation affinity, resulting in increased monomer levels.
For this project, I won’t be depleting TDP-43 directly, but I will be inducing a similar effect. A few members of our lab figured out which amino acids allow for dimer formation. Mutating these sites can create reduced dimer and increased monomer levels. For this experiment, I am using the N-terminal mutant Thao Nguyen created, which involves the mutation of three amino acids (not to be disclosed for confidentiality reasons) in the N-terminal of TMEM106B.
Specifically, I will be conducting wet lab experiments using a neuronal cell model to determine how TMEM106B’s protein interaction partners differ between TMEM106B monomers and dimers. This research will provide new molecular insights into the mechanisms by which FTD/ALS develop that may contribute to the advancement of future therapeutics.
Originally, this project was going to examine how changes in polyA site usage (alternative polyadenylation) in the 3’-UTR upon TDP-43 knockdown could impact transcript protein interaction partners or protein interaction partners in TMEM106B, but we have decided to slightly change the direction to focus on monomer vs. dimer formation. However, I hope to also investigate protein interaction partners in TMEM106B length changes upon APA using the same techniques if time permits.
That’s it, and I’m looking forward to updating you guys with my progress throughout the next few months!
Glossary
FTD: frontotemporal dementia, a neurodegenerative disease.
ALS: amyotrophic lateral sclerosis, a neurodegenerative disease.
TDP-43: normally performs necessary functions in the nucleus like RNA regulation, maladaptively mislocalized in neurodegenerative diseases like FTD and ALS which leads to loss-of-function in the nucleus (can't perform necessary functions) and gain-of-function in the cytoplasm (aggregates to form harmful stress granules).
APA: alternative polyadenylation, an RNA regulation process that describes the use of different polyA (polyadenylation) sites on mRNA. When APA occurs, mRNA is cleaved at a different polyA site (to which the polyA tail is then added), leading to a lengthened or shortened transcript.
3'-UTR: 3' untranslated region, a gene expression regulatory region (not translated into protein) that starts after the stop codon, regulating mRNA stability and translation. All polyA sites in TMEM106B are in this region.
TMEM106B: the gene I will be studying APA in, due to its common dysfunction in frontotemporal dementia.
N-mut: a mutant construct our lab created, involving mutations in Cys 61, Cys 64, Thr 63 in the N-terminal of TMEM106B.
Week 1: Transfection
Hi everyone, welcome back to my blog!
As a small recap, a few members of our lab figured out which amino acids allow for dimer formation. Mutating these sites can create reduced dimer and increased monomer levels. For this experiment, I am using the N-terminal mutant (which I will be referring to as 'N-mut') Thao Nguyen created, which involves the mutation of three amino acids in the N-terminal of TMEM106B.
I used the pmaxGFP, pmirGlo-TMEM106B-GFP N-mut, and pmirGlo-TMEM106B-GFP WT (wild-type) constructs. These constructs include the introduction of GFP (green fluorescent protein) so I can later check to make sure transfection worked. N-mut is the mutation that disrupts dimer formation as mentioned above, and WT encodes the normal TMEM106B protein. PmaxGFP is the empty vector control with only GFP (no TMEM106B) so I can later differentiate which proteins are sticking to GFP vs. TMEM106B.
Transfection
This week, I began transient transfection to introduce plasmids with this amino acid mutation into HEK (human embryonic kidney) cells. Transfection is the process of introducing DNA into cells through non-viral means. Because cells are surrounded by a lipid bilayer, many substances don’t easily pass through. To introduce my DNA into the cells, I added lipofectamine 3000 (lipo 3000), creating a cationic (allowing it to be attracted to the negative DNA) lipid encapsulation to allow it to enter the lipid bilayer. P3000 is another required reagent for lipid formation.
Although I will combine the DNA, P3000, and lipo 3000, the protocol recommends separation with two tubes for each construct in the beginning. Tube 1 for each construct will include 5µg of DNA and 40µL P3000 suspended in opti-MEM reduced serum medium, adding up to 300µL total. Tube 2 will be the same for each construct, with 40µL of lipo3000 suspended in opti-MEM medium, also adding up to 300µL total.
To streamline the process, I started by pipetting 260µL of optiMEM into each tube. For the lipo3000 tubes, I then pipetted 40µL lipo3000 into each tube.
Because the plasmids have different concentrations and thus result in different volumes being needed, the DNA + P3000 tube will just be slightly less straightforward to combine. Before I began, I had to calculate how much of each plasmid I needed for our desired 5µg of material based on their given concentrations. Because I will introduce each plasmid to three wells in my HEK cell plates to ensure consistency, I also tripled each value to determine how much total of each plasmid I need. The calculations are as follows:
pmaxGFP (1µg/µL): 5µg/(1µg/µL) = 5µL
Tripled: 15µL
N-mut (0.551µg/µL): 5µg/(0.551µg/µL) = 9.07µL
Tripled: 27.22µL
WT (0.469µg/µL): 5µg/(0.469µg/µL) = 10.66µL
Tripled: 31.98µL
For each DNA + P3000 tube, I sucked out the amount of optiMEM equivalent to the amount of plasmid we will be adding (15µL, 27.22µL, 31.98µL, respectively). Then, I added in the proper amount of DNA. Finally, I pipetted 40µL of P3000 into each sample.
Now that I was finished with creating all the DNA + P3000 and lipo3000 tubes, I combined them by pipetting all 300µL of each lipo tube into the DNA tubes.
Next, I grabbed three HEK cell plates (6 wells each) and labeled each with the plasmid I was introducing. I divided the volumes of each of my three tubes by 6 to pipette each plasmid evenly into the six wells of their respective plate.
The next day, I took all the cells to the fluorescence room and imaged them to check to see if my transfection worked. I am able to detect in which cells transfection was successful because they would express GFP. Below are my results:
We see that transfection worked! The glow is much more intense for the GFP construct because it only expresses GFP, but there is still a glow for N-mut and WT as well. I also included an image of the N-mut construct with the GFP-expressing cells overlaid on the bright field image (in red, showing all the cells) to show how many of the cells transfection was successful in.
Next, I took the plates and sucked out all the opti-MEM media. I washed each well with 1mL PBS then removed the PBS. I added 500µL NP40 buffer to each well to lyse the cells and solubilize their proteins. I waited around 15 minutes, then scraped off any remaining cells stuck on the bottom with a cell scraper. To ensure consistency, I combined every 2 wells in each 6-well plate into 1 Eppendorf tube, resulting in 3 tubes per construct (GFP 1, GFP 2, GFP 3, N-mut 1, N-mut 2, etc.). Finally, I took these tubes to the sonicator, a machine that makes an unpleasant noise but (on the bright side) shears the DNA. This allows for the proteins to get released into the solution and ensures all cells are lysed properly.
Transformation
Before I did transfection, I also conducted E. coli transformation to amplify the amount of plasmid I have to keep my supply of plasmids for future use.
I searched up which antibiotic resistance each construct possesses and found that kanamycin for pmaxGFP and ampicillin for the others. With this information in mind, I grabbed 1 kanamycin agar plate and 2 ampicillin agar plates. Because I only want to collect E. coli in which transformation was successful, introducing an antibiotic will kill all E. coli cells in which the transformation (introduction of the plasmid) was not successful.
I first combined my competent cells (E. coli) and each plasmid sample into three separate tubes. Next, I heat-shocked them to promote uptake by the E. coli and put them back on ice quickly after. After adding SOC media to the heat-shocked cells to allow them to better “recover,” I added some of each tube into their respective agar plates (kanamycin for pmaxGFP and ampicillin for the others). I added some magnetic beads and shook the plates to distribute the transformed E. coli solution throughout the plates. After I finished, I incubated them overnight.
Next week, I am planning to do some BCA and immunoprecipitation - see you all in my next blog post! :)
Week 2: BCA
This week, I conducted a BCA to measure the concentrations of my samples to determine how much volume of each sample I need to add for next week's immunoprecipitation.
To avoid thawing and freezing the original sample I obtained from transfection last week too much, I further took out 10µL into tiny tubes to use for my BCA.
First, I made 5x dilutions from the tiny tubes. I grabbed a new set of tiny tubes, pipetting 40µL of NP40 buffer and 10µL of sample into each. I used NP40 buffer because it’s lighter than harsh detergents like RIPA buffer, allowing it to preserve the protein interaction partners we’re aiming to find.
In my 96-well plate, I included each material in triples to ensure consistency. I pipetted 10µL of BSA standard proteins with known concentrations in ng/µL (125, 250, 500, 750, 1000, 1500, 2000) into the first 7x3 rows, which will allow me to find the equation relating concentration to absorbance, thus enabling me to find the concentrations of my own samples. I then pipetted 10µL NP40 buffer into the 8th row. Finally, I pipetted 10µL of my samples each into the following rows in triples.
Then, I prepared a working solution, which involves a blue liquid and a clear liquid, to be added in a 1:50 ratio. I multiplied the number of wells I used (51) by 200µL (I want to use ~200µL in each well), getting 10200µL. Then, I rounded up to 11000µL and divided this number by 51 to determine how much blue liquid I need (215µL). Accordingly, I pipetted 215µL of blue liquid and 11000µL-215µL=10785µL of clear liquid into a tube. Then, I dumped this entire mixture into a reservoir and pipetted 180µL into each well (200µL is ideal, but I did 180µL to ensure I don’t run out of material). Finally, I placed it into an incubator. After half an hour passed, the samples turned various shades of purple or green:
I analyzed the BCA using a machine called the spectrophotometer, which gives me the absorbances of every well in my plate. I downloaded the data Excel and began analyzing my results.
After creating a line of best fit of each standard concentration (for each, I averaged the absorbances of the triples), I obtained an equation:
Using this equation, I solved for the concentrations of each of my samples. I then multiplied each value by 5 to account for the dilution I did at the very beginning (when I combined 10µL sample and 40µL NP40 buffer). I divided each number by 1000 to convert from ng/µL (the units of the standards) to µg/µL.
My final product for BCA will be made of my samples of equal mass (500µg) and NP40 buffer. I divided 500µg by each concentration to determine how much volume of each sample I need. Originally, I was aiming for a total of 500µL per solution, but I didn’t have enough NP40 buffer, so I changed the total volume of the solution for each into the volume of the largest sample (375µL).
After pipetting the appropriate amounts into 9 tubes (labeled “i” or immunoprecipitation), I pipetted 15µL of each into 9 new tubes. This will be my “starting material,” which I will be using later.
I didn’t get to start immunoprecipitation this week, so I will be doing so next week. See you guys then!
Week 3: Immunoprecipitation
In a process called immunoprecipitation, adding magnetic beads to my samples will bind to the GFP tags of my desired proteins, which I can then isolate by attracting the complex to the magnetic rack the samples are placed within. This week, I did immunoprecipitation and prepared a gel for mass spectrometry.
I first took out the 375ml sample + NP40 buffer solutions I made from last week (labeled “i”). After I finished creating more NP40 buffer because I ran out, I began immunoprecipitation.
Immunoprecipitation
I grabbed a solution of GFP-trap magnetic beads from the fridge and took out 100µL to add to a new tube, which I then placed on a magnetic rack. The magnetic beads quickly separated from the storage buffer, allowing me to suck out the undesired storage buffer.
After washing the beads two times with 1mL NP40 buffer (note: I was supposed to wash three times), I added 100µL NP40 buffer. Then, I pipetted 10µL of the solution into each of my samples in the “i” tubes and placed them onto the rotator.
Putting each solution back on the magnetic rack after waiting for 4 hours, the beads quickly separated from the NP40 buffer. I sucked out the NP40 buffer and kept them in 9 new tubes labeled “ft” (flow-through). I then washed the solution 3 times with NP40 buffer again.
I spun the tubes down to remove residual wash buffer before resuspending the beads of each sample in a 20µL gel loading dye solution containing reducing reagent (DTT). To make this solution, I rounded up to 10 tubes to ensure I had enough material and multiplied by 20µL for a total of 200µL. As my gel loading dye stock concentration is 4x and the recommended concentration is 2x, I made this solution with 100µL of gel loading dye. I then added 20µL of 10x concentrated reducing reagent and milli-Q water for the remaining 80µL.
Finally, I boiled each sample for 10 minutes and put them back onto the freezer. Boiling the samples allows for my desired proteins to detach from the magnetic beads and get suspended in the gel loading dye.
The next day, I loaded and ran an SDS-Page gel with my samples still in the magnetic rack, making sure to avoid taking in magnetic beads. I put my finished gel in coomassie blue stain, then put it onto the rotator for around 30 minutes. When it was done, I placed it in 25% methanol.
When I checked my gel the next day, a few of the bands were less concentrated, indicating that I might not have added the same amount of beads to each sample.
Thus, I took another day to redo immunoprecipitation (beginning from remaking “i” tubes based on last week’s calculations). Unfortunately, I forgot to make new flow-through tubes for these samples, but hopefully it’s not a problem.
I’ll be running my gel with my re-done immunoprecipitation samples next week! Hopefully the beads for each sample turn out equal this time.
Week 4: Troubleshooting and Verification
Hi guys, welcome back to my blog! This week, I mostly focused on redoing immunoprecipitation, the steps of which are explained in detail in my previous post, and verifying the existence of TMEM106B in my samples.
Troubleshooting
After I prepared a gel from my 2nd immunoprecipitation of last week and soaked it in coomassie blue, I discovered that the gel bands were barely visible, thus having to redo my gel again (now onto my third gel). While preparing this gel, I suspect that a few mistakes I made such as breaking the walls between the gel wells and putting in a Kim wipe (which is supposed to absorb excess coomassie blue) during the coomassie blue staining stage rather than the methanol washing stage led to my bands being improperly stained.
I prepared my third gel with the unused samples that I loaded my wells with, learning from my mistakes and making sure to be very careful at every step. When the gel was ready, I placed it into coommassie blue and put it on the shaker, then replaced coommassie blue with 25% methanol after an hour and put in a Kim wipe. I replaced the 25% methanol and Kim wipe after another hour and left it on the shaker overnight.
When I checked my gel the next day, the bands still turned out a little hard to discern from the gel which didn’t destain fully, but they were still visible and looked consistent.
Before sending my gel to the mass spectrometry facility in Australia through my mentor Steph, I removed my gel from the methanol it was soaking in. I placed it on a glass panel and cut each band out with a scalpel, then moved each band into a new labeled tube: finally ready for sending!
However, since I used all of my loading samples, I had to prepare new ones for verification, thus beginning immunoprecipitation round 3. It took a while, but eventually all the samples were newly prepared and ready for verification!
Verification
To verify the presence of TMEM106B in my samples, I’m running western blots for both my ip samples and my starting material samples (see Week 2 blog). As a recap, I collected 15µL of each sample/NP40 solution after BCA for my starting material. This way, in case I don’t see TMEM106B in my ip gel, I can determine if it’s due to unsuccessful immunoprecipitation or if it was never there in the first place.
From a quick Google search, I expect to see the empty GFP vector constructs to be 27kB, while the GFP-TMEM constructs with an additional TMEM106B would be heavier.
I split the starting material into two separate sets, one with reducing reagent and one without. Typically, gel samples are run with reducing reagent; however, reducing reagents break disulfide bonds, changing both N-mut and WT TMEM106B into mostly monomers. Thus, I expect my starting material gel to have no noticeable difference between the N-mut and WT bands.
This week, I finished running the gel for SM and ip. However, before I left, I accidentally broke the ip gel while cracking it open, so I will have to redo it next week. As I had to leave before I was able to finish preparing my gel, my mentor Steph helped me run transfer buffer through my gel onto a membrane and left the membrane in blocking buffer in the cold room shaker overnight.
Week 5: More Verification
This week, I continued running more western blots as verification, while my mass spectrometry sample was getting processed in Australia. I started the week off by redoing my IP gel.
Western Blots (More Verification)
As I had limited amounts left of each IP sample, this time I loaded each well with 5µL instead of 10µL. When the gel finished running, I assembled a sandwich made of a stack of transfer paper, transfer membrane, my gel, and another stack of transfer paper (in that order, bottom-up) within a cassette. Before adding each layer, I soaked them in transfer buffer for 1-2 minutes to prevent them from drying.
After locking the cassette, I drained out the remaining buffer and placed it into the Trans-Blot Turbo machine. After 7 minutes, I carefully peeled off the membrane and placed it into another container. Finally, I added blocking buffer to the container and placed it on the shaker overnight, next to the SM membrane. The proteins that blocking buffer contains stick to the background of the membrane, allowing primary antibodies to bind more accurately to our desired proteins instead of background noise.
On the next day, I took out both membranes and dumped the blocking buffer. To save the amount of material I need to use for all subsequent steps, I moved my ip membrane and SM membrane to be in the same box, making sure to note which membrane is which. The materials we will be introducing will be able to permeate both membranes.
Next, it was time to add the primary antibodies. For my western blots, I wanted to identify two proteins: my TMEM106B and GAPDH, a loading control. The concentration (intensity) of GAPDH allows me to determine how even each sample is to each other. Primary antibodies are either derived from rabbits or mice. As secondary antibodies (which bind to primary antibodies) are species-specific, I used a solution that had both rabbit and mouse antibodies to differentiate TMEM106B and GAPDH. I then left the membranes on the shaker for 1 hour.
After 1 hour, I created my secondary antibodies mixture. I added 10mL blocking buffer and 0.5µL of each secondary antibody. The rabbit secondary antibody emits green fluorescence in later imaging, while the mouse secondary antibody binds to the GAPDH primary antibody and emits red fluorescence. I left the membranes on the shaker for an hour again.
When finished, I took out the membranes and washed them with TBST buffer 3 times. In between each wash, I left them on the shaker for 10 minutes. Finally, I covered the membranes with TBST again to prevent them from drying and took them to an imaging room. I then labeled my images using PowerPoint. The following were my results:
As we can see, the SM image is very clear. However, I can’t really see anything on my ip membrane, so I added primary antibody again and left it on the shaker overnight in the cold room for its primary antibodies to hopefully have more time to bind to the proteins. This is likely due to the fact that I didn’t load enough material, and many samples didn’t flow to the bottom of the well due to the lack of enough glycerol from loading dye.
The next day, I redid the remaining steps again and obtained the following image:
The image was still very faint, so I dramatically turned up the intensity, thus resulting in lots of background noise as well. However, the presence of TMEM106B monomers do seem to be visible. TMEM106B dimers might also be there for WT, but it’s hard to be certain.
The day after, I decided to run another gel. In this gel, I loaded one sample of each triple for SM (this time without reducing reagent, as SM with reducing reagent is running low) and ip, a loading control (beads suspended in only loading dye, not pulling down anything compared to my ip samples), and two wells of just loading dye to fill up my remaining wells.
I ran the same western blot procedure, except this time with more (and only) GFP for the secondary antibodies (2µL instead of 0.5µL). I then obtained the following image:
(Note: because I had washed the well too harshly, some of the ladder leaked over to its adjacent rows)
As the ip was very light, I decided to redo the gel again next week in hopes of yielding better results.
Week 6: ...and More Verification
This week, I started by running yet another gel. This gel is very similar to the last gel I ran last week, except the two samples of only gel loading dye occupy the last two wells.
This gel used 8µL of the same SM as I used last week, but I used my ip samples from the very first mass spectrometry gel I ever attempted (see Week 3). My loading control was made of 7µL (this was a mistake and should’ve been 8µL), washed with NP40 buffer 3 times, and 10µL gel loading dye, 2µL reducing reagent, and 8µL milli-Q water (then finally boiled at 95˚C for 10 minutes).
After my gel finished running, I set up another membrane stack. However, when I loaded the cassette containing my membrane stack into the Trans-Blot Turbo machine, I forgot to press run and my membrane looked suspiciously bare, but I kept it in blocking buffer overnight anyway. My suspicions that the proteins didn’t successfully transfer onto the membrane were confirmed when I soaked it in Ponceau S for 10 minutes, a red stain that exposes proteins on your membrane, and didn’t see any bands.
The next day, I redid the entire process again. This time, the transfer was successful!
When I repeated the remaining steps of adding antibodies, washing to get rid of excess antibodies and reagents, and imaging, I obtained a nice (and finally presentable) image!
In this image, it is very clear in the SM columns that WT contains both monomers and dimers, while N-mut contains mostly monomers. The dimer band isn't as intense in the IP columns' WT, but the trend still looks pretty clear as well for the most part.
Additionally, my mass spectrometry results came back this week. In the following weeks, I will be analyzing the results I have obtained.
See you all again for next week’s blog!
Week 7: Data Analysis, Transformation, Validation, & Cell Culture
Last week, my mass spectrometry results finally arrived in the form of an Excel sheet with over 2000 proteins!
I first set some filters on the spreadsheet. I made it so that they only include proteins that are increased or decreased 2-fold while comparing between N-mut and WT; this way, I only have proteins that actually see a change between the two constructs. To do so, I entered “‘greater than or equal to 2’ or ‘less than or equal to 0.5’” in the column comparing each protein’s abundance in N-mut to WT (“NmutGFP/WTGFP”). I then created two separate sheets for when NmutGFP/WTGFP >/= 2 and </= 0.5 (ordered from highest difference to lowest difference), along with separate sheets for NmutGFP/EVGFP (empty vector GFP) and WTGFP/EVGFP.
Interestingly, my TMEM106B levels are much higher in WT than N-mut, suggesting that I didn’t add equal amounts as I should have.
Right off the bat, Steph noticed cathepsin D, a protein involved in lysosomal protein cleavage, to be in my list. Steph then ordered a cathepsin D antibody, so I can verify whether cathepsin D really is in my samples (using my western blot membrane from last week). When I later compared my list to some pre-existing proteomics studies, the suggested association of this protein was further strengthened by Takahashi, H. et al. (2023), who also found this protein in their paper Lysosomal TMEM106B Interacts with Galactosylceramidase to Regulate Myelin Lipid Metabolism. We also found cathepsin A under the name “lysosomal protective protein” in my list.
To begin making sense of the rest of my proteins, I skimmed through and highlighted all of the proteins that had familiar or intriguing names. While highlighting, I noticed some interesting proteins: other TMEM proteins (TMED9, TMEM33, TMEM109, and of course, TMEM106B), cell cycle regulation/tumor-related proteins (CDK5, BCCIP, TPD52), proteins related to ubiquitin (NDUFAF3, UBE2S, ARIH1, USP5, UBE2D3, OTUB1), and those involved in autism, especially the mTOR pathway (LAMTOR2, CSNK2A1, FKBP1A, FMR1, ATP6V1E1, SEC13).
I then searched up lysosomal proteins and highlighted all proteins in my list that are in the lysosome. These proteins are particularly interesting because TMEM106B has been found to have cleavages in its C-terminal, which is the segment of the protein within the lysosome. A notable one that is supported by Lübke, T. (2009) includes LAMP2.
When the cathepsin D antibody arrived, I did the antibody steps on my western blot membrane again. This time, the primary antibody solution was made of 12mL blocking buffer and 4µL cathepsin D mouse antibody, and the secondary antibody solution was made of 12mL blocking buffer and 0.5µL goat anti-mouse secondary antibody. The cathepsin D antibody that we bought didn’t have a recommended western blot dilution ratio listed on its website, so I had to look through some papers’ methodologies to seek advice. However, they were not very helpful either, although I did find one paper that used a 1:3000 dilution for a different company’s cathepsin D antibody in combination with a 1:100 dilution for the one I have. 1:100 seemed too large though, so I just decided to try a 1:3000 dilution (4µL) for now, a recommended ratio for ThermoFisher’s cathepsin D antibody product.
When I checked the image results, cathepsin wasn’t visible, likely because 4µL was not enough to detect enough primary antibodies. I tried doing a 1:100 dilution this time, adding 120µL - 4µL = 116µL cathepsin D antibody to the same primary antibody solution I had used earlier. I added my newly concentrated solution to the membrane and left it overnight on the cold room shaker.
The next day, I finished the remaining antibody and wash steps and took it to the imager again. Although I couldn’t see any new signs, when I turned up the intensity very high in the red channel (which shows mouse antibodies), I could see a faint signal. When comparing it to the turned-up red channel intensity of the original membrane without cathepsin D antibody, the signals were only present in the new membrane, making it a promising sign!
Transformation
The rest of the week, I mostly spent more time on data analysis and began to work on my final project presentation, but I also began transformation to redo the entire process once again in hopes of obtaining the perfect western blot with definitive results about the cathepsin antibodies.
Cell Culture
Then on Friday, I also split some cells to prepare new cells for my next goal, to knock down the cathepsin proteins in HEK293T cells (if I find definitive cathepsin signals from the western blot) and see what happens as a result of this depletion.
Week 8: More Validation
This week, I began to restart the entire process again to acquire more material for further validation.
Inoculation
To prepare for a midi prep, I took my three samples from last week’s transformation and started inoculation. I grabbed 3 new falcon tubes and labeled them GFP, WT, and N-mut. As Steph had pre-prepared liquid LB with kanamycin (the antibiotic that the GFP construct is resistant to), I added 35mL of this solution into the GFP falcon.
To make liquid LB with the ampicillin antibiotic, I added 35mL LB broth to the WT and N-mut falcons. To add the ampicillin antibiotic in a 1:1000 dilution, I added 35µL to each falcon.
Next, I opened the LB agar plates one at a time and inoculated a single colony from each using a pipette tip. I dropped each tip into its respective falcon and swirled the falcon before pouring its contents into a new 250mL beaker. After covering each beaker with aluminum foil, I left them in the shaking incubator overnight.
Plasmid Midi Prep
The next day, I took my flasks from the incubator, and they were cloudy - a sign that there has been bacterial growth! I then conducted a midi prep to extract and purify my amplified plasmids. When I was finished, I left them in the -20˚C freezer for later use.
Contamination (!)
When I checked on my HEK293T cell confluency (see last week’s blog), I discovered an unfortunate situation — my cells were contaminated! Sadly, we had to bleach the cells and dispose of them.
Hopefully, we’ll get a chance to split more healthy cells next week when Steph’s HEK293T cells are confluent enough for more splitting.
More Validation
For the rest of the week, I did some more validation. As our cathepsin A antibody finally arrived, I added it to the membrane I’ve been using for the past few weeks (the one with both SM and IP). I obtained the image below:
That’s it for this week. Hope to see you all again next week!
Week 9: More Validation (Cont.)
This week, I ran another (likely my final) western blot. Because I have more material from the very first immunoprecipitation I did (the one I labeled “i,” see Week 3’s blog), I checked the first gel I ran for mass spectrometry (which failed because it was uneven) and used the 3rd replicate of each construct as it was the most consistent.
I ran my western with SM (without reducing reagent) and i, using the 3rd replicate of each. This time, instead of verifying for TMEM106B first, I focused on cathepsin D (sc-13148, mouse 1:500) and cathepsin A (Proteintech 15020-1-AP, rabbit 1:1000).
Because I had to leave early, I asked another member of the lab to help complete the western blot transfer onto a membrane, to which he graciously agreed. He left my membrane in EveryBlot blocking buffer in the cold room overnight.
The next day, I took my membrane from the cold room and created/added my cathepsin D solution, then left it on the shaker for 1 hour. After washing with TBST for 30 minutes, I added my cathepsin A solution onto the membrane and left it on the shaker for another hour, before washing with TBST for another 30 minutes.
I added my secondary antibody solution made of 2µL mouse and rabbit antibody each and EveryBlot blocking buffer and left it on the shaker. Finally, I repeated the remaining steps of a western (washing).
When I went to image my membrane, the bands were so faint that I had to dramatically turn up the intensity to see them, until the background noise was huge. I separated the scan into three separate images:
Before leaving for the day, I left the membrane in the cold room.
As there is way too much background noise, not enough signal, streaks in some lanes, and lots of non-specific binding (indicated by the presence of multiple bands in each lane), I had to change something about the way the antibodies bind to the membrane.
The next day, I stripped the membrane of its antibodies using stripping buffer. To do so, I washed the membrane with TBST, added the stripping buffer, incubated for 30min, washed with TBST again, then blocked with 5% milk in TBST for an hour. This allows me to start the antibody binding process from scratch.
Then, I repeated the remaining steps. However, this time I used milk (1% in TBST for each antibody solution) instead of EveryBlot blocking buffer for each step. Milk is an incredibly effective blocking buffer as it’s packed with proteins, so hopefully it’s more effective at blocking background noise than EveryBlot was. I also left it in a new primary antibody solution I made (using milk) in the cold room overnight this time to allow for the antibodies to have more time to bind to the proteins.
The results were as follows:
As the images still aren’t very clear, Steph speculates that the antibodies just aren’t very good and advised me to focus on data analysis and my project final product and presentation.
That’s it for this week! See you guys again next week for my (nearing final) blog!
Week 10: Final Product
We’re nearing the end of this project!
As I’ve finished the wet lab portion of my project last week, this week I’ve been working on my final product, a mini paper. Although I’ve spent the whole week working on this paper, a lot of this involved shortening all the blogs I’ve written over the course of this project. Because this wouldn’t be new information, below is a little look into just the introduction I’ve rewritten, now that I’m nearing the completion of my project and have refined its direction.
Introduction
Frontotemporal dementia (FTD) is a neurodegenerative disease caused by frontotemporal lobar degeneration (FTLD) (1). Around half of FTLD patients possess TAR DNA binding protein 43 (TDP-43) inclusions, making up the largest category, FTLD-TDP (2-3).
Transmembrane protein 106B (TMEM106B) is a protein whose genetic variations have been found to be a key driver of FTLD-TDP (4-8). TMEM106B resides within the membranes of late endosomes and lysosomes and plays important roles in regulating transportation and maturation within the lysosome (9).
The TMEM106B protein is made of three domains: the N-terminal suspended in the cell’s cytoplasm, the transmembrane domain, and the C-terminal suspended within the lysosome (10-11). Cryo-electron microscopy points to pathological C-terminal fragments forming amyloid fibrils in patients with FTLD-TDP (12-15). These insoluble fragments contribute to decreased TDP-43 function (1).
Although TMEM106B filaments are a normal part of aging, genetic variations lead to differential expressions of FTLD-TDP (1). Patients who possess the TMEM106B rs3173615 protective allele experience a slowdown of FTLD-TDP progression, along with postmortem samples showing little aggregation, in contrast to increased levels for those with the risk allele (1). Those with the protective allele saw increased levels of TMEM106B dimers, suggesting the role of dimers in limiting fibril formation (1).
[...] (this section is cut out for confidentiality)
To further understand the impact of TMEM106B monomerization on FTLD-TDP disease progression, this study will investigate the differences between the proteomes of TMEM106B monomers and dimers.
References
Arai https://www.sciencedirect.com/science/article/pii/S0006291X06023187?via%3Dihub
Gallagher https://pmc.ncbi.nlm.nih.gov/articles/PMC4003885/
Blitterswijk https://pmc.ncbi.nlm.nih.gov/articles/PMC3944829/
Van Deerlin https://pmc.ncbi.nlm.nih.gov/articles/PMC2828525/
Jiao https://molecularneurodegeneration.biomedcentral.com/articles/10.1186/s13024-023-00644-1
Lang https://www.jbc.org/article/S0021-9258(20)49998-3/fulltext
Schweighauser https://pmc.ncbi.nlm.nih.gov/articles/PMC9095482/