Culturing Primary Drosophila Neurons

Though there has been great progress with understanding actin and microtubule dynamics, and their relationship to cell motility, the scientific community is still working to try to better define these interactions, and their associated proteins, that allow this movement to happen. Many neurodevelopmental disorders and degenerative diseases are linked to genes associated with the cytoskeleton (Prokop et. al., 2013), making impaired motility of neurons a possible cause. Therefore, further information on growth and motility dynamics could help lead potential treatments for these disorders.

Neurons are understood to develop an axon, their outwardly-signaling appendage, by the elongation of a neurite via microtubule recruitment. The growth cone at the tip of this developing axon is actin-rich, and forms focal adhesions with the substrate to facilitate movement and network development (Prokop et. al., 2013), making the growth cone the leading edge in neurons.  Interestingly, there have been some indications that these growth cones may move at different rates on some substrates than others. Because of the substrate-dependency lead, it may be possible to utilize different substrates to characterize some of the proteins present in the extracellular matrix that contribute to efficient movement of neurons. For my thesis project I’m attempting to look at growth cone dynamics on two substrates: extracellular matrix (ECM) and concanavalin A (ConA, a plant lectin often used to plate cells) using neurons extracted from Drosophila 3rd instar larvae.

Drosophila lend themselves as an excellent model for examining the role of proteins in neuronal movement. Not only are the proteins affecting actin and microtubule dynamics well-conserved, making the findings generalizable across species, but the genetics are simplified in drosophila as well, with little redundancy in the genome. This attribute makes it so that a single knockout is capable of almost completely eliminating the function of a protein, whereas in other animals, such as mice, the knockout of multiple genes may be required to silence a protein due to redundancy (Prokop et. al., 2013). Additionally, existing knowledge about their genome makes it so that we can easily and time-efficiently make genetically altered strains for experimentation. This makes knockdown studies easy and efficacious, and desirable crosses quick to generate. Their neurons have been found to grow well in cultures, even forming networks and displaying similar electrical properties to in-vivo functioning neurons when in proper media (Küppers-Munther et. al., 2004). This allows easy visualization and analysis of neuronal movement, further contributing to their experimental value, which is why drosophila were chosen by our lab for these studies.

In a typical experiment, I extract 6-10 brains from the drosophila larvae and apply liberase to break apart the cell-cell contacts. These neurons are then plated in rich cell media on either an ECM- or a ConA-coated glass slide in a dish and allowed to attach to the bottom of the slide. After the neurons have attached, I visualize their movement by taking videos under a microscope. Additional visualization of proteins can be achieved with staining.

Sources:

Küppers-Munther, B., Letzkus, J. J., Lüer, K., Technau, G., Schmidt, H., & Prokop, A. (2004). A new culturing strategy optimises Drosophila primary cell cultures for structural and functional analyses. Dev Biol, 269(2), 459-478.

Prokop, A., Beaven, R., Qu, Y., & Sánchez-Soriano, N. (2013). Using fly genetics to dissect the cytoskeletal machinery of neurons during axonal growth and maintenance. J Cell Sci, 126(Pt 11), 2331-2341.

Actin Microtubule Crosslinking

Studies have shown that the cytoskeletal elements in the cell (actin, microtubules and intermediate filaments) engage in extensive crosstalk. This crosstalk is an important part of the regulation of the cytoskeleton, as well as a number of other biological processes. For this blog post, I will be focusing on actin-microtubule cross linking, since that is most relevant to my thesis research. Before I get into specifics about how this is related to my thesis, I’m going to give a brief, general overview of actin microtubule crosstalk and the various roles it can play within a cell.

Actin is a highly conserved protein in cells that switches between G-actin and filamentous F-actin. Actin is one of the most abundant proteins in eukaryotic cells and plays an important role in muscle contraction, cell signalling and regulation of cell shape. Microtubules are dynamic, polar and are made up of alpha & beta tubulin. The tubulin polymerizes to form microtubule filaments.  Key to microtubules ability to perform their functions in the cell is dynamic instability and polarity. Dynamic instability is characterized by periods of rapid growth followed by periods of depolymerization. This allows microtubules to rapidly alter their configuration in order to fit the needs of the cell. Microtubules and actin play important roles in cell division, cell migration and other important cellular processes.

Actin Microtubule crosstalk is mainly defined through the physical mechanisms by which it occurs. This means that usually what happens is a physical linkage between parts of actin and parts of microtubules that lead to stabilization or nucleation etc. One of the main forms of actin microtubule crosstalk is crosslinking. This occurs when proteins link microtubules to actin. This linkage is enabled by large protein complexes that can also interact with microtubule plus end binding proteins. This linkage connects the plus ends of microtubules to actin bundles which can result in a redirection of microtubule growth.

One of the proteins that plays a role in actin microtubule cross linking is Short Stop (Shot). Short stop is a spectraplakin that has been shown to bind microtubules & actin filaments and has also been localized in growth cones. In my thesis, I’m looking at microtubules in Drosophila melanogaster neuroblasts, specifically at what happens when you knock down shot thus inhibiting crosslinking. I’m studying the effect of knocking down this crosslinking protein on microtubule dynamics in 3 different parts of the neuron.

 

Works Cited

 

Applewhite, D.A., Grode, K.D., Keller, D., Zadeh, A.D., Slep, K.C., and Rogers, S.L. (2010). The Spectraplakin Short Stop Is an Actin–Microtubule Cross-Linker That Contributes to Organization of the Microtubule Network. Molecular Biology of the Cell 21, 1714–1724.

Dogterom, M., and Koenderink, G.H. (2018). Actin–microtubule crosstalk in cell biology. Nature Reviews Molecular Cell Biology.

Sanchez-Soriano, N., Travis, M., Dajas-Bailador, F., Goncalves-Pimentel, C., Whitmarsh, A.J., and Prokop, A. (2009). Mouse ACF7 and Drosophila Short stop modulate filopodia formation and microtubule organisation during neuronal growth. Journal of Cell Science 122, 2534–2542.

qRT-PCR to verify RNAi knockdown

As discussed in a recent blog post, RNAi is a common technique used in the Applewhite lab to observe the effects of a silenced gene. When preparing for RNAi, it is common practice to run a sample of the dsRNA template on a gel to make sure the resulting band is the same size as the target. However, is this sufficient practice to conclude your following results are due to the knockdown of the gene? Many publishers would say no. It is possible that the exogenous dsRNA was not in appropriate concentration, or was not an effective target to cleave the specific mRNA sequences. Real time quantitative polymerase chain reaction (qRT-PCR) is a supplementary method used to verify the successful knockdown of the gene of interest.

Quantitative PCR (qPCR) is accomplished by extracting endogenous RNA from your cells treated with RNAi, reverse transcribing the RNA to DNA, designing primers to amplify the gene of interest, and using intercalating dyes, such as SYBR Green, which bind to the DNA and fluoresce with greater intensity as the concentration of the target sequence increases. Important to note, in the presence of off-target dsDNA, sequence-specific probes can be used which rely on FRET for detection, and fluoresce only when the DNA polymerase separates the quencher from the emitter. These sequence-specific probes include Taqman, Molecular Beacons, and Scorpions, although require more complex and expensive implementations (1).

The real time element is essential to determining initial DNA template concentration. Since qPCR only measures the end concentration of target sequence, there is no way to calculate an initial concentration. qRT-PCR however, measures template concentration at an exponential stage of replication, which allows for calculation of an initial starting concentration. This in turn, enables analysis of initial gene expression, and if minimal, verification of RNAi success (2).

Figure 1. Relative qRT-PCR and qPCR measurements of target concentration in respect to duration of PCR.

Sources Cited

  1. The Basics: RT-PCR.Thermo Fisher Scientific – USAvailable at: https://www.thermofisher.com/us/en/home/references/ambion-tech-support/rtpcr-analysis/general-articles/rt–pcr-the-basics.html. (Accessed: 23rd October 2018)
  2. Bansal, R.et al.Quantitative RT-PCR Gene Evaluation and RNA Interference in the Brown Marmorated Stink Bug.Plos One11,(2016).

 

Mander’s Coefficient

One aspect of my thesis is exploring co-localization of Split Discs with other proteins in drosophila cells. In order to do this, not only does wet lab work need to be accomplished, but mathematical analysis (in this case using Mander’s coefficient).

Fluorescence microscopy does not have the ability to see whether or not two molecules are directly interacting. However, by looking to see if they co-localize in the cell, it can be determined whether they interact with the same complexes in the cell. The limit for fluorescence microscopy is the resolution of the images produced. Because of this, small numbers of puncta are not sufficient for determining whether or not the experimental molecules are actually co-localized. Multiple puncta from different regions within the cell must be used in analysis so the data is not limited to overlapping puncta which are a result of organelles that are close in proximity to one another.

In order to quantitatively determine the correlation of co-localization in the cell, mathematical analysis of the data is employed. For my thesis, I am employing Mander’s Overlap Coefficient (MOC) for this analysis because it does not require distinguishing fluorescence as being the result of a fluorescent protein or background noise. MOC is able to do this because it only compares the co-occurrence of fluorescence among pixels. MOC = ∑i(Ri×Gi) / √(∑iR2i×∑iG2i) where Ri and Gi are the average level of grey from the red and green fluorescence respectively (Manders et al., 1993). MOC has a range of 0 – 1 and Ri and Gi have a range of -1 – +1. The limitation to this equation is that the ratio of values can result in ambiguous numbers. Therefore, the numerator and denominator can be split up in such a way to account for the ambiguity.  From this we get two coefficients: M1 (fraction of red fluorescence in areas with green fluorescence) and M2 (fraction of green fluorescence in areas with red fluorescence)  (Manders et al., 1993). M1 = (∑iRi,colocal) / ∑iRi where Ri,colocal = Ri if Gi > 0 and Ri,colocal = 0 if Gi = 0 and M2 = (∑iGi,colocal) / ∑iGi where Gi,colocal = Gi if Ri > 0 and Gi,colocal = 0 if Ri = 0 (Manders et al., 1993). The larger MOC, M1, and M2 are the stronger the evidence for co-localization of the proteins within a cell. In my thesis, MOC, M1, and M2, will be gathered for each cell to determine whether or not Split Discs are co-localizing with other specific proteins.

 

References:

 

Dunn, K. W., Kamocka, M. M., & McDonald, J. H. (2011). A practical guide to evaluating colocalization in biological microscopy. American Journal of Physiology – Cell Physiology, 300(4), C723–C742. http://doi.org/10.1152/ajpcell.00462.2010

Manders, E. M., Verbeek, F. J. & Aten, J. A. (1993). Measurement of co‐localization of objects in dual‐colour confocal images. Journal of Microscopy, 169, 375-382. doi:10.1111/j.1365-2818.1993.tb03313.x

BioID of SPECC1L

BirA is biotin protein ligase in E. coli that selectively adds biotin to a subunit of acetyl-CoA carboxylase. Roux et al. created a mutant BirA that isn’t selective to its native substrate and will instead biotinylate any proteins that are close-by. This allows for identification of protein-protein interactions in eukaryotic cells by creating a fusion protein of BirA and a protein of interest that will biotinylate proximal proteins, which can then be captured and identified. Protein identification utilizes the strong association of biotin and streptavidin (or avidin) to capture proteins that have been biotinylated by BirA, followed by mass spectroscopy to identify biotinylated proteins.

 

Figure 1. Schematic of BioID assay from Roux et al. 2012.

I am investigating the protein interactions of an actin-microtubule crosslinking protein coded by the gene SPECC1L (with an ortholog known as “split discs” in Drosophila). Mutations in SPECC1L have been identified in a number of diverse cases of orofacial cleft, and thus knowledge of protein-protein interactions by the product of SPECC1L is particularly important to understanding the developmental basis of this set of conditions.

To this end, I am using a BirA-split discs construct to investigate what proteins split discs interacts with in vivo in Drosophila. This construct will ideally biotinylate proteins that split discs typically interacts with, without excessive non-target biotinylation and without affecting the behavior or split discs. I will then capture biotinylated proteins using the biotin-streptavidin interaction and identify using mass spectrometry.

 

References:

Roux KJ, Kim DI, Raida M, Burke B. A promiscuous biotin ligase fusion protein identifies proximal and interacting proteins in mammalian cells. The Journal of Cell Biology. 2012;196(6):801.

CG and Flapwing: Knockouts, Knockdowns, and RNA Interference

When attempting to understand a protein’s function in a cell, the effects of removing that protein can be very telling. As such, knockdowns and knockouts, techniques which remove a targeted protein from acting in the cell, are widely used to identify how a cell behaves without the functions performed by said target protein.

 

A knockout is an irreversible procedure that removes the target protein from the cell permanently, often by editing the genome itself to either deactivate or directly remove the target protein’s coding sequence. This can be achieved by selective breeding when dealing with whole organisms, but when working with cell cultures, knockouts are typically accomplished by means such as the gene-editing kit TALEN or, in recent years, by use of the tool CRISPR-cas9.
A knockdown, on the other hand, is a repeated procedure which removes the target protein from the cell temporarily. By treating cells on a regular basis, the target protein is disrupted from its usual function, and the cells can be observed without its influence. If the treatment ceases, the target protein will no longer be disrupted and the cell will return to normal function, if mildly worse for wear. The distinction between these two procedures can be likened to ending a fight (knockout) as opposed to temporarily ‘gaining advantage’ or suspending the fight (knockdown).

 

In our experiment, we have performed a RNA interference (RNAi) knockdown of our target proteins flw and CG. This is achieved first by the transformation of genomic DNA (gDNA) that codes for the target protein into double stranded RNA (dsRNA), and then by simply exposing cultured cells to the dsRNA on a sustained and regular basis.
The reason this apparently simple method (which requires more pipetting and tube shuffling than that short sentence might imply) works and removes the target protein from action is due to the cell’s own inherent defensive mechanisms. When exposed to free floating dsRNA in solution, some dsRNA is naturally taken up into the cell, where it is recognized as foreign and chopped to pieces. Ironically, this causes the fragmented dsRNA to bind to messenger RNA (mRNA) already in the cell that matches its sequence, whereupon the entire dsRNA-fragment-mRNA amalgamation is recognized as foreign and chopped to pieces.

 

Figure 1. dsRNA suspended in cell media is taken into a cell, recognized as a foreign component, and cut into pieces by the appropriate enzymes. The cut pieces of dsRNA attach to pieces of mRNA naturally present in the cell, which are subsequently tagged for sheering due to their binding with a foreign component.

 

When the dsRNA is in sufficient concentration, this defensive response results in the cell being unable to translate the appropriate mRNA into the target protein, since it is instead destroying that mRNA as fast as it can. Once depleted of the target protein, cells can be treated, fixed, stained, or any combination thereof, and the effects of the knockdown can be observed to extrapolate the target protein’s function.

Team Force: Data Analysis Techniques

In the previous post we described how the data is collected using Traction Force Microscopy (TFM). The process of imaging outputs several “movies,” which display the cells exerting forces and moving the beads embedded in the compliant gel matrix.

The data analysis algorithm is threefold: 1) tracking, 2) low-pass filtering, and 3) calculating traction stresses.

To calculate the displacements of the beads, a reference image of the beads on a plain compliant matrix is compared to an image of the beads on a compliant matrix with the cell on top of it which then pulls on the substrate near its edges. Using an array containing the tracked particle data for each frame of the movie, the displacements are extrapolated with stochastic drift taken into account. These data are then output onto an XY grid at each time interval.

Then, a low pass-filter is added to remove the high-frequency noise from the displacement data.

Next, the displacement data is correlated to the traction stresses through an algorithm derived by Style et. al. (2014) along with the elasticity theory, which states that the properties of the compliant matrix such as thickness, stiffness, and compressibility must be taken into account when considering traction stresses, which are continuous distributions of forces (Abidi, 2016). Modeling the gel as a “spring,” we can use Hooke’s Law, F=-kx, where k is the elasticity constant, x is displacement, and F is the force.

Finally, once the traction stresses have been computed, we will overlay a plot of the displacement and traction stress vectors on top of an image of the cell, as shown in Figure 1 (Abidi, 2016).

Figure 1: A force vector field calculated by Abidi (2016) using example data from Style et. al. (2014)

 

References

Abrar A. Abidi. Quantifying cellular mechanotransduction in morphogenesis and cancer. Reed College, 2016.

Robert W Style, Rotislav Boltyanskiy, Guy K German, Callen Hyland, Christopher W MacMinn, Aaron F Mertz, Larry A Wilen, Ye Xu, and Eric R Dufresne. Traction force microscopy in physics and biology. Soft Matter, 10(23):4047-4055, 2014.

 

 

CG and Flapwing: creating clones using synthetic vectors

One way we intend to examine the role of our target proteins, CG and Flapwing (flw) is by RNA interference knockdown (RNAi, in which the target proteins are inhibited in live cells, those cells exposed to the apical constriction signaling ligand, Fog, and the cells fixed so their response may be quantified. But in addition to knockdown, we also plan to measure target overexpression and localization. To these ends, we will use recombinant DNA techniques to produce clones of our target proteins’ coding sequences, in the form of a synthetic vector.

 

The process of producing a vector applicable to the proteins of interest first requires that template DNA of both targets are amplified by polymerase chain reaction (PCR). Once the DNA has been scaled up, it is precipitated and resuspended in water before being “digested” by two specialized restriction enzymes that sever the DNA at specific locations in sequences artificially inserted into the coding gene for the target proteins.

Because the cleavage sites on any two antiparallel sites cut by the same restriction enzyme will be complimentary, it is possible to place complimentary cut sites on both our insert and a cloning vector (pMT/V5-His A), thus allowing the cloning vector’s cut ends to match to the cut ends of our insert and for the two to bind. The process of combining vector and insert and encouraging them to bind and circularize is known as ligation.

Once ligation is complete and the insert DNA has been successfully added to the vector, the product is introduced to a bacterial culture and allowed to propagate before the culture is spun down by centrifuge and the DNA is isolated, for further use in increasing the expression of our target proteins in cell culture.

Due to difficulties with the cloning procedures, we are still currently attempting to successfully force the vector to take up the insert and vectorize the genes of our target proteins, which is necessary to introduce this increased protein expression into cell cultures. We have as such added CIP to the PMT His-A digestions, which prevents the vector from re-forming with itself and requires the binding of the insert for the DNA to circularize, and are expecting better results from our ligations in the future.

 

References:

 

  1. Molecular Cell Biology. 4th edition. Lodish H, Berk A, Zipursky SL, et al. New York: W.H. Freeman; 2000
  2. Thermo Fisher Scientific (2018). Traditional Cloning Basics | Thermo Fisher Scientific – US.

PARF Analysis

PARF analysis or Permeabilization Activated Reduction in Fluorescence analysis allows researchers to describe the rate of dissociation of a fluorescently tagged protein of interest from intracellular structures. I will primarily be spending most of my time conducting PARF analysis to determine the effect of depleted SPECC1L on focal adhesions.

In our experiments, we transfect, or introduce, fluorescently tagged vinculin, which is one protein known to be found in focal adhesions. We visualize the focal adhesions using TIRF microscopy, and take control videos for each condition. Primarily, however, we are interested in the rate of dissociation of the fluorescently tagged vinculin from focal adhesions. This rate of dissociation corresponds to the size and relative strength of the focal adhesions. To cause this dissociation, we add digitonin to the cell media as we are filming to induce permeabilization of the cell membrane. To see more, look at the bottom of the post for a supplemental video!

This permeabilization results in an unbalanced gradient of fluorescent vinculin, with a high concentration within the cell, and a low concentration in the surrounding media. This gradient favors the dissociation of the vinculin into the external media. We measure the loss of fluorescence for around two minutes and then can fit this to an exponential decay. Using both a positive and negative control that causes larger and smaller focal adhesions, we can determine the effect of depleted SPECC1L on a cell’s focal adhesions.

Supplementary Material:

PARF Blog Post Attachment

Supplemental Figure 1. This technique can be applied to various other proteins of interest. In the above video, we permeabilize these S2R+ cells expressing both fluorescent Naus GFP and fluorescent mCherry cortactin with digitonin after 20 seconds. We can then visualize the loss of fluorescent cortactin as it dissociates from the cell into the surrounding media.

Note: To view the video, you have to download it! If you have a Mac, click on the link while pressing Ctrl, and download the file.

Citations:

  1.  Singh PP, Hawthorne JL, Quintero OA. Permeabilization Activated Reduction in Fluorescence (PARF): a novel method to measure kinetics of protein interactions with intracellular structures. Cytoskeleton (Hoboken). 2016 June ; 73(6): 271–285. doi:10.1002/cm.21306.

Team Force

Our team will use a physical approach to investigate the role of SPECC1L (Split Discs) in cellular contractility by quantifying force expression through Traction Force Microscopy (TFM).

In TFM, cells are adhered to compliant gel matrices with fluorescent beads. The traction forces are found as shown in Fig. 1

Figure 1: Measurement of cellular contraction in cells (Jacobs et al., 2012)

The movement of the fluorescent beads is tracked using the microscope, and along with the known physical characteristics of the substrate such as thickness and stiffness, we can extrapolate the force vectors between the cell and gel matrix.

We are using gene inhibition using dsRNA induced gene silencing (RNAi) to target proteins such as Spaghetti Squash (Sqh), Myosin Binding Subunit (Mbs), and our protein of interest, SPECC1L. We will compare the force expression of Split Discs against that of Sqh depleted cells, which will cause the inactivation of non-muscle Myosin II (NMII) and hypocontractile cells, as well as Mbs depleted cells, which will result in the absence of dephosphorylation of RLC, causing an open NMII and hypercontractility (Abidi, 2016).

References:

Abrar A. Abidi. Quantifying cellular mechanotransduction in morphogenesis and cancer. Reed College, 2016.

Christopher R. Jacobs, Hayden Huang, and Ronald Y Kwon. Introduction to Cell Mechanics and Mechanobiology. Garland Science, 2012.