Cas Staples
Actively utilizing the 3D DNA conformation to engineer gene expression requires a tool that is highly specific while still allowing for targeting any location within the genome. Recently, CRISPR/Cas-based systems that manipulate three-dimensional DNA conformation to regulate cellular processes have been developed. However, these systems are often complex and have a limited range of applications. Here we present the design and application of fusion guide RNAs (fgRNAs) and fused Cas-nucleases to create a simple, adjustable system for the precise programming of DNA-DNA contacts. We successfully demonstrated the use of these Cas staples and their application in gene expression control by induced proximity of DNA strands. Taken together, our results establish Cas staples as an innovative and powerful tool for remodeling the three-dimensional landscape of the genome.
Introduction
The CRISPR/Cas System as a Gene Editing Tool
In 2012, Jinek et al. described the use of the Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas system, to induce DNA double-strand breaks. Since their seminal work, gene editing with CRISPR/Cas has come a long way as exemplified by a multitude of applications in all areas of the life sciences and even first CRISPR-based gene therapies (Sheridan, 2023). At their core the CRISPR/Cas gene editors are constituted by a ribonucleoprotein complex consisting of a Cas nuclease and one or two RNA molecules responsible for guiding the nuclease to a specific genomic site. The most widely applied CRISPR nuclease is Cas9 from Streptococcus pyogenes. It binds a CRISPR RNA (crRNA) and a transactivating CRISPR RNA (tracrRNA) that can be combined into a single guide RNA (sgRNA) (see fig. 1A) (Pacesa et al., 2024). The sgRNA includes a scaffold required for its binding to the Cas nuclease and an interchangeable 20 nucleotide (nt) spacer sequence that defines the DNA target via complementary base pairing. Once Cas9 binds to a matching DNA strand, it efficiently cleaves the target DNA (Cong et al., 2013). Furthermore, a specific three nucleotide sequence (NGG) on the 3’ end in the targeted DNA is required for binding and cleavage. This is referred to as the protospacer adjacent motif (PAM) (Sternberg et al., 2014).
Figure 1: The CRISPR/Cas system. A and B, schematic structure of Cas9 and Cas12a with their sgRNA/crRNA, sitting on a DNA strand with the PAM. The spacer sequence forms base pairings with the dsDNA. In case of Cas9 the spacer is located at the 5’ prime end, for Cas12a at the 3’ end of the gRNA. The scaffold of the gRNA forms a specific secondary structure enabling it to bind to the Cas protein. The cut sites by the cleaving domains, RuvC and HNH, are symbolized by the scissors (adapted from Pacesa et al. (2024)).
Over the following years additional Cas proteins with different functional properties have been discovered, including Cas12a from Acidaminococcus sp. (AsCas12a) and Moraxella bovoculi (MbCas12a)(Zetsche et al., 2015). In contrast to Cas9, the gRNAs of Cas12a carry have a 5’-scaffold (see fig. 1B). Similarly, the PAM (TTTN) is also on the 5’ side of the spacer (Pacesa et al., 2024). Important to our work, Cas12a is capable of processing arrays of multiple consecutive crRNA repeats into individual crRNAs/gRNAs enabling the expression of multiple gRNAs from a single expression cassette (Paul and Montoya, 2020). The introduction of specific mutations allows the generation of catalytically dead dCas variants (Koonin et al., 2023) (Kleinstiver et al., 2019). By fusing dCas proteins to transactivator domains and targeting them to an endogenous promoter region via a specific gRNA, they can be harnessed for programmable transcriptional activation or “CRISPR activation” (CRISPRa) (Kampmann, 2017).
Strategies to Program DNA Interactions
The three-dimensional conformation of genomes is crucial for the correct interpretation of the genetic code and the phenotypic plasticity of cells. Interacting genetic elements often occur in close proximity in cis but it was shown that long-range interactions of genomic loci separated by thousands of bases or even located on different chromosomes are similarly important (Cramer, 2019). Here, the three-dimensional (3D) conformation of the genome comes into play and directly influences biological functions (Lieberman-Aiden et al., 2009)(Dixon et al., 2012). Being able to artificially influence DNA interactions would allow us to study and program the 3D genome organization and to study and potentially cure pathophysiological rearrangements. One complex example to achieve this, makes use of the CRISPR/Cas system in the form of light-activated dynamic looping for endogenous gene expression control (LADL) by Kim et al. (2019). In this system, two dCas9 proteins bind to specific endogenous loci through their respective sgRNAs. The dCas9 are fused to photoreceptors that cluster upon activation by blue light resulting in the inducible bridging of two genomic loci. (see fig. 2).
Figure 2: Light-activated dynamic looping for endogenous gene expression control. The Cas proteins bind to the DNA via the sgRNA. They are fused to CIBN, which, upon exposure to blue light, forms a heterodimer with CRY2. CRY2 also forms oligomers with other CRY2 proteins, resulting in the bridging between two different Cas9 protein complexes. This induces the looping of DNA segments (adapted from Kim et al. (2019)).
Aim of this subproject
Although existing systems such as LADL principally allow the manipulation of the 3D DNA conformation, they
require several components and several gRNAs, resulting in a very complex system. Besides its complexity
LADL relies on the unspecific clustering of dCas9 rendering multiplexing impossible. Related less complex
systems, that are
based on smaller proteins like zinc finger domains, remain difficult to adapt to any genomic locus of
choice (Kim and Kini, 2017).
Here we present CRISPR/Cas based protein staples, that build and expand related systems for gene editing.
We create fusion guide RNAs (fgRNAs) that are composed of a Cas12a and Cas9 gRNA, connected via their
spacers (Kweon et al., 2017). This system solely
requires the two Cas proteins dCas9 and dCas12a and an fgRNA to bring two elements
close together. This allows for precise, adjustable targeting while keeping the system simple in general.
By introducing several subsequent fgRNAs, this may also allow for multiplexing. By harnessing the Cas12a’s
ability to process chimeras of Cas9 and Cas12a into individual units we can decide exactly which genes are
brought into proximity and which not (Gonatopoulos-Pournatzis et al., 2020).
The first part of this project focuses on establishing functional fgRNAs. We then continue to show that
these fgRNAs can be used in combination with Cas12a and Cas9 to form a complex that brings two separate
DNA loci into proximity.
Results
To successfully induce proximity of two DNA strands we had to connect two different DNA binding elements. We selected SpyCas9 and MbCas12a as DNA-binders due to their programmability. To physically connect both proteins, we decided to link their gRNAs. In contrast to fusing the Cas proteins, this comes with the advantage that one could easily multiplex the system, while still guaranteeing that specific pairs of genomic loci are connected. Specifically the 3'-end of the Cas12a gRNA was fused to the 5'-end of the Cas9 gRNA. Via this approach the two spacer sequences are fused directly, ensuring a minimal distance between the two DNA strands. The design also facilitates interchangeability, as the central portion of the fgRNA construct, which includes both spacers (for Cas9 and Cas12a) together with an optional linker, can be easily replaced in one cloning step. The scaffold components remain integrated within the backbone of the construct. To implement this compact composition and ensure an efficient cloning procedure, we engineered an entry vector for the fgRNAs . We inserted a gene block encoding ccdbB between the scaffolds of Cas12a and Cas9 guide RNA (gRNA). SapI cut sites were incorporated between the scaffolds and the ccdB gene. Therefore, all fgRNAs can be created by a simple Golden Gate Assembly when adding the spacer sequence as annealed oligonucleotides to the entry vector (see fig. 3). CcdB ensures bacteria transformed with the unchanged plasmid to die due to the toxicity of ccdB. This concept was proven by our own clonings throughout the project resulting in a very high success rate for the transformations. Picking one colony for sequencing was sufficient in all cases but one.
Figure 3: Construction process of fgRNAs using the entry vector. The ccdB gene can be cut out using SapI in the Golden Gate assembly. By inserting oligonucleotides with the desired spacer sequences and matching overhangs, the complete fgRNA can be expressed. Due to the cytotoxic nature of ccdB, only cells with the oligonucleotides as inserts survive.
Editing endogenous loci with fgRNAs
To develop and test functional fgRNA Cas staples we determined two essential applications of the fgRNAs to establish first. These are the utilization for multiplex genome editing and the implementation in CRISPRa (see fig. 4) (Kweon et al., 2017).
Figure 4: Applications of the Fusion Guide RNA. Fusion Guide RNAs can be used for multiplex genome editing by guidingactive Cas12a and Cas9 to two distinct loci. Similarly, fgRNAs allow for CRISPRa, by guiding the Cas9-VP64 transcriptional activator towards a target locus (adapted from Kweon et al. (2017)).
To prove that our fusion gRNAs still result in active ribonucleoproteins, a series of different
fgRNAs were created, each carrying spacers specific to the VEGFA and FANCF genes.HEK293-T cells
were transfected with the Cas protein and gRNA constructs.
The editing rate was tested 72h after transfection via a T7 endonuclease I assay.
AsCas12a and SpCas9 were used. The AsCas12a spacer targets
VEGFA, while the SpCas9 spacer targets FANCF. The samples included standard single gRNAs
with the corresponding Cas protein, the fgRNA with only one of the two Cas proteins and
the fgRNA with both Cas proteins simultaneously (see fig. 5).
The sgRNAs allowed for the highest editing rates for both genes (45% for VEGFA and 15% for FANCF),
while the editing rates for FANCF were consistently lower in all experiments. Importantly,
targeting FANCF with fgRNAs resulted in noticeable editing of about 10%, with just the SpCas9 and
both Cas proteins in the sample. For VEGFA, the AsCas12a only sample resulted
in approximately 20% editing rate in combination with the fgRNA, while adding both Cas proteins
led to approximately 40%. These initial results confirmed our engineering approach proving
efficient genome editing with fgRNAs.
Figure 5: FgRNAs Enable Efficient Editing of Endogenous Loci.The editing rates were determined 72h after transfection via T7EI assay. Editing % was determined by measuring band intensities; Editing % = 100 x (1 – (1- cleaved band/uncleaved band)) 1/2. The schematic at the top shows the composition of the fgRNA. Below each spacer is the targeted gene. The symbols below indicate which parts are included in each sample.
Fusion Guide RNAs Allow for Editing Rates With Variant Cas Orthologs
To further evaluate the capabilities of the fgRNAs, we tested them in combination with
different Cas12a orthologs. After some initial testing, we decided on using MbCas12a together with
SpCas9.
Additionally, to test if the differences in editing rates from the preliminary assay resulted from
the targeted loci or the different Cas orthologs, the spacers were tested in both arrangements.
Once with Cas12a targeting FANCF and SpCas9 targeting VEGFA and once vice versa. To better assess
the impact that the utilization of a fgRNA has on the editing rates, the sgRNAs were tested
separately and in one sample.
Having the sgRNA with single Cas proteins in the same sample resulted in no clear
difference in the editing rates (see fig. 6A and fig. 6B). The fusion of the gRNAs resulted in
a lower editing rate overall. While the editing for VEGFA stayed at about 20% in all
cases, the editing for FANCF dropped significantly. When targeting the same gene under the same
conditions, the editing rates for MbCas12a were overall lower than the ones from SpCas9.
Figure 6: Fusion gRNA Editing Rates In Combination with MbCas12a. A and B, the editing rates were determined 72h after transfection via T7EI assay. Editing % was determined by measuring band intensities; Editing % = 100 x (1 – (1- cleaved band/uncleaved band)) 1/2. The schematic at the top shows the composition of the fgRNA. Below each spacer is the targeted gene. The symbols below indicate which parts are included in each sample. A and B display both orientations of the two spacers for VEGFA and FANCF.
The Inclusion of a Linker Does Not Lower Editing Rates
To further assess the effect of the genomic locus on the editing rate, we included CCR5 as an additional gene target. For this assay, a fgRNA with a 20 nt long linker was included between the two spacers. The editing rate for VEGFA was again relatively consistent throughout the samples (see fig. 7). For CCR5, the editing rate with sgRNAs was approximately the same at about 30%. However, it dropped below 10% for the fgRNA. The addition of the 20 nt linker had no effect on the editing rates compared to no linker.
Figure 7: Fusion gRNA Editing Rates for Multiplexing CCR5 and VEGFA The editing rates were determined 72h after transfection via T7EI assay. Editing % was determined by measuring band intensities; Editing % = 100 x (1 – (1- cleaved band/uncleaved band)) 1/2. The schematic at the top shows the composition of the fgRNA. Below each spacer is the targeted gene. The symbols below indicate which parts are included in each sample. Cas12a targets VEGFA and Cas9 targets CCR5.
Using fgRNAs for CRISPRa
To establish the foundation for their use as protein scaffolds, we identified the next step as demonstrating the use of fgRNAs for CRISPR activation. For this, we intend to recruit the transcriptional activator VP64 to a firefly luciferase gene to induce expression. The VP64 protein is attached to the catalytically inactive Cas9 protein, which is then guided by gRNAs to the luciferase gene. The gRNAs target a TetO sequence, which is positioned in front of the luciferase gene in multiple repeats. The firefly luciferase activity was then quantified as photon counts and normalized against Renilla luciferase, which is expressed on a separate plasmid under an ubiquitous promoter. In two biological replicates we saw similar Relative luciferase activity with fgRNA as a guide compared to a sgRNA (see fig. 8). For further insight into the engineering behind these findings, we recommend to take a look at the engineering cycle fgRNA iteration 4 and 5.
Figure 8: CRISPRa Induced Luciferase Expression for sgRNAs and fgRNAs. Firefly luciferase activity was measured 48h after transfection. Normalized against ubiquitously expressed Renilla luciferase. The tetO repeats were targeted by Cas9-VP64, once with a sgRNA and once with a fgRNA that had a non-targeting sequence for the Cas12a spacer. The schematic at the top shows the composition of the fgRNA. Below each spacer is the targeted gene. The symbols below indicate which parts are included in each sample.
Stapling Two DNA Strands Together Using fgRNAs
After showing the general capability of the fgRNA to work for editing and for CRISPR activation,
the next step was to use it to staple two DNA loci together, and thereby induce proximity between
two separate functional elements. For this, an enhancer plasmid and a reporter plasmid was used.
The reporter plasmid has firefly luciferase behind several repeats of a Cas9 targeted sequence.
The enhancer plasmid has a Gal4 binding site behind several repeats of a Cas12a targeted sequence.
By introducing a fgRNA staple and a Gal4-VP64, expression of the luciferase is induced (see fig. 9A).
Different linker lengths were tested. Cells were again normalized against ubiquitous renilla
expression. Further information on our learnings from this assay can be found in the Cas staple
engineering cycle iteration 2.
Using no linker between the two spacers showed similar relative luciferase activity to the
baseline control (see fig. 9B). An extension of the linker from 20 nt up to 40 nt resulted in an
increasingly higher expression of the reporter gene.
Figure 9: Applying Fusion Guide RNAs for Cas staples. A, schematic overview of the assay. An enhancer plasmid and a reporter plasmid are brought into proximity by a fgRNA Cas staple complex binding both plasmids. Target sequences were included in multiple repeats prior to the functional elements. Firefly luciferase serves as the reporter gene, the enhancer is constituted by multiple Gal4 repeats that are bound by a Gal4-VP64 fusion. B, results of using a fgRNA Cas staple for trans activation of firefly luciferase. Firefly luciferase activity was measured 48h after transfection. Normalized against ubiquitously expressed Renilla luciferase. Statistical significance was calculated with ordinary One-way ANOVA with Dunn's method for multiple comparisons (*p < 0.05; **p < 0.01; ***p < 0.001; mean +/- SD). The assay included sgRNAs and fgRNAs with linker lengths from 0 nt to 40 nt.
To further improve the efficacy, we introduced a GSG-linker between the Cas proteins. Similar to
the initial tests for the fgRNAs, the capability of our fusion Cas constructs was tested by
assessing the editing rates via a T7EI assay. For this, the same target sequences as before were
used, namely FANCF and VEGFA in both configurations. We included biological duplicates in this
assay.
The fusion Cas proteins allowed for editing in general, with single gRNAs and fgRNAs (see fig.
10A and 10B). The editing rate with Cas9 was higher overall. Especially for the fgRNA fusion Cas
combinations, the Cas12a editing rates were significantly lower, dropping to about 1%. At the same
time, targeting VEGFA resulted in a higher editing efficiency than FANCF, as one can see in our
fgRNA engineering cycle iteration 3.
Figure 10: Editing rates for fusion guide RNAs with fusion Cas proteins. A and B, the editing rates were determined 72h after transfection via T7EI assay. Editing % was determined by measuring band intensities; Editing % = 100 x (1 – (1- cleaved band/uncleaved band)) 1/2. The schematic at the top shows the composition of the fgRNA. Below each spacer is the targeted gene. The symbols below indicate which parts are included in each sample. Cas proteins linked by a dash ("–") were fused to each other. Biological replicates are marked as individual dots.
To further investigate the characteristics of the fusion Cas system, CCR5 was again included as a different target site, as well as a 20 nt linker between the two spacers. In this case, while the combination of fusion Cas proteins with sgRNAs allowed for a high editing rate of 15% to 25% at both target sites, VEGFA in combination with Cas12a was much more consistent at around 20% than CCR5 with Cas9 at about 2% (see fig. 11). The inclusion of a linker had no significant impact on the editing rates with a fgRNA.
Figure 11: Targeting VEGFA and CCR5 with fusion Cas proteins and fgRNAs. The editing rates were determined 72h after transfection via T7EI assay. Editing % was determined by measuring band intensities; Editing % = 100 x (1 – (1- cleaved band/uncleaved band)) 1/2. The schematic at the top shows the composition of the fgRNA. Below each spacer is the targeted gene. The symbols below indicate which parts are included in each sample. Cas proteins linked by a dash ("–") were fused to each other.
Continuing the procedure in a similar manner as for the fgRNAs, we focused on inducing proximity between genetic loci next. The same assay was used, with one enhancer plasmid and one reporter plasmid. Though less distinct than the results for using just fgRNAs, the fusion Cas proteins can be used to increase expression levels of the reporter firefly luciferase (see fig. 13). While using sgRNAs results in similar relative luciferase activity as for the negative control between 0.1 and 0.2., using a fgRNA with a 20 to 30 nt linker consistently resulted in activities at 0.25. Fusion guide RNAs without a linker and with a 40 nt linker had on average about the same activity, but with a higher spread over the biological replicates. Further learnings from this assay and how we want to continue in the future is layed out in our Cas staples engineering cycle iteration 2.
Figure 12: Results of Implementing Fusion Cas Proteins in Trans Activation of a Reporter Firefly luciferase activity was measured 48h after transfection. Normalized against ubiquitously expressed Renilla luciferase. Statistical significance was calculated with ordinary One-way ANOVA with Dunn's method for multiple comparisons (*p < 0.05; **p < 0.01; ***p < 0.001; mean +/- SD). Fusion Cas proteins were paired with sgRNAs and fgRNAs with linker lengths from 0 nt to 40 nt.
Discussion
The use of fgRNAs for multiplex gene editing with Cas9 and Cas12a was shown to be effective. Though the
fgRNAs have proven to allow for similar editing rates compared to sgRNAs in some cases, a number of
factors have been identified that have a higher impact on the efficiency of fgRNAs, including the targeted
genomic locus and the Cas ortholog. The editing rate varies considerably for different genes. In the
various assays conducted in this subproject, VEGFA showed a relatively consistent and high editing
rate. The editing rate of FANCF was observed to be fluctuating and lower in most cases.
This is likely due to differences in chromatin accessibility, which allows the Cas proteins
to reach some parts of the DNA more effectively than others (Klemm et al., 2019).
Cas9 and Cas12a appear to not only have a varying editing rate overall, but also show
different responses to the fusion of the gRNAs or the Cas proteins themselves. Comparing
editing rates on individual genes indicates an overall increased performance of Cas9 with
fgRNAs.
One potential explanation for this observation might be a higher tolerance of Cas9 to
modifications made to the gRNA. In contrast, the addition of a linker appears to have
no impact on the editing rates. A reason for that might be that either both Cas proteins
are not able to bind together in general, or that 20 nt are not enough space between the
spacers to fit both Cas proteins.
While the fusion Cas protein constructs also worked in combination with fgRNAs, their overall perfomance
was better in presence of sgRNAs.
Having two individual connections in the Cas staple complex, might result in a more rigid assembly that
would require precise coordination of the discrete linkers. To achieve this, a vast range of protein and
gRNA linkers would need to be tested in different combinations, allowing for a better assessment on the
exact effect they have on the system and how these linkers would need to be combined to allow for an
effective forming of the Cas staple complex.
In comparison to the results presented by Kweon et al., Cas9 was also observed to
have
a higher editing rate in general. However, the difference in editing rates for different
genes was not significant and the results show the editing rates for FANCF were in fact
higher. To further assess the actual difference the targeted gene makes, a large scale screen
of multiple different genes that are targeted under the same conditions would be necessary.
The main objective of this project was to establish fgRNA based Cas staples as an effective tool to bring
genetic loci of choice into proximity. We first showed the fgRNAs ability to bind DNA in conjunction with
a dead Cas protein, by applying CRISPRa.
In the subsequent assays, we were able to show that our fusion Cas construct is able to bring and hold two
genomic loci in proximity in a way that allows for gene activation. By this we not only confirm that
changes in the 3D genome structure can determine whether a gene is activated, but also show that our Cas
staple system is capable of exactly that.
Though, compared to simple CRISPRa, the fold changes of
expression are way lower, this may allow for way more precise manipulation of gene expression. By
expanding the system to endogenous loci and enhancers, it could easily be linked to complex regulatory
systems within the cell, permitting a precisely engineered gene expression for the locus of choice.
Introducing fusion Cas proteins into the Cas staple system, showed first promising results, though not
significant enough to consider it as a clear increase in gene expression. This again shows the requirement
of further assays to understand the characteristics of fusion Cas proteins, especially in the context of
binding DNA.
The use of CRISPRa with fgRNA aligns with the results previously published in this regard, also showing an
increase in gene expression through the recruitment of an activator (Kweon et al.,
2017).
Changing the 3D structure to induce expression with fgRNA staples is to our knowledge a completely new
approach. In a similar way, this has been done with the LADL system (Kim et al.,
2019). Other approaches that use a more complex, modified version of Cas9 also showed that gene expression
can be altered by chemically inducing rearrangements on a 3D-level (Morgan et al.,
2017). These publications also show, that hijacking genomic loci for this procedure is feasible.
Outlook
To further prove our Cas staples to be applicable in shaping the human genome, a few new assays were
designed to be conducted in the future.
We are already in the process of designing constructs for the next experiments, which would focus on using
human genome enhancers to increase expression of target genes.
This extension of the system would further show how the introduction of Cas staples changing the
arrangement of DNA is enough to influence expression levels at will.
In a very recent publication the usage of so-called double guide RNAs (dgRNAs) for a similar application
was proposed (Yang et al., 2024). These dgRNA consist of two subsequent Cas9 gRNAs
rather than a Cas12a and a Cas9 gRNA.
While this publication only showed application in bacterial context, we want to properly assess and
compare both systems in the eukaryotic environment. We are currently in the cloning phase of dgRNAs that
are usable in HEK293-T cells and plan to perform the same assays as we did with our fgRNA system to
thoroughly compare these to ways of using gRNA fusions as the basis of DNA stapling.
On the one hand, the fgRNA system would be able to solve complex systems via multiplexing different target
sites, for which we already have an experiment running, confirming this. On the other hand, the dgRNA
needs only one Cas ortholog to form the staple complex, lowering the amount of protein coding DNA required
to be introduced into the host cell. In this may lie the possibility of making it easier to package in
delivery
systems like adeno-associated viruses and thereby enabling the use in gene therapy.
Having both of these systems available in our toolbox, further increases the range of applications that we
can provide to researchers.
Furthermore, we plan on improving the fusion Cas staple, to get similar expression levels as seen for the
fgRNAs. First thoughts on new approaches can be read about in our Cas staples engineering cycle iteration
2. This would again include the screening of different protein linkers combined with different fgRNAs. The
establishment of a stable protein linker is the basis of making it responsive to outside stimuli like
certain chemicals or proteases. In the context of staple extension we show the successful cleavage of a
peptide linker by cathepsin B, allowing for the functionalization of a Cas staple construct. We would thereby enable our system to be adaptable to different conditions, that can
be specific to certain types of cancers through the likes of tumor micro environments or overexpression of
distinct proteins (Anderson & Simon, 2020).
Staple Functionalization
Cathepsin B is a lysosomal protease present in the cytosol of various cancer types. We overexpressed cathepsin B in HEK293T cells to investigate its ability to cleave different peptide linkers using a fluorescence readout assay. We successfully demonstrated that the GFLG linker was efficiently cleaved by cathepsin B in vivo when cells were treated with doxorubicin. Furthermore, we showed that wild-type cathepsin B matured into its active forms under these conditions. Together, these findings enable the functionalization of our PICasSO toolbox for a wide range of therapeutic and synthetic biology applications.
Introduction
Cathepsin B is a cysteine protease typically located in lysosomes or secreted outside the cell, where it degrades proteins of the extracellular matrix. It plays a critical role in apoptosis and is significantly overexpressed in various cancer types, including breast and colorectal cancer (Ruan et al., 2015). The overexpression of cathepsin B is associated with tumor invasion and metastasis. Various stimuli, such as ischemia, bile acids and TNFα, can induce cathepsin B-mediated apoptosis. In this process, lysosomes release large amounts of cathepsin B into the cytosol, where it cleaves anti-apoptotic factors like Bcl-2 and XIAP, leading to an increase in apoptotic proteases, such as caspase-3 (Bien et al., 2010).
The significance of cathepsin B in cancer progression is well-documented, with studies showing elevated cathepsin B levels in cancerous tissues compared with non-cancerous tissues (Ruan et al., 2015). Research has shown a connection between elevated levels of cathepsin B and enhanced angiogenesis, invasion and metastasis (Ruan et al., 2015). Given its important role in tumor progression, cathepsin B is considered a potential therapeutic target (Ruan et al., 2015) or prodrug-activating enzyme (Zhong et al., 2013). Proteolytic cleavage of pro-biologics allows for the precise temporal and spatial regulation of biopharmaceutical activity in therapeutic strategies (Bleuez et al., 2022).
Aim of this subproject
We explore the potential of our PICasSO platform approach for therapeutic applications, we designed protein-based DNA staples that are responsive to the overexpression of cathepsin B in cancerous tissues. Furthermore, We demonstrate doxorubicin-dependent cathepsin B cleavage of one out of five documented linkers (Jin et al., 2022; Shim et al., 2022; Wang et al., 2024) in HEK293T cells.
Additionally, we develop a construct consisting of one dead Cas9 (dCas9) connected to two SV40 nuclear localization sequences and two caged intein fragments (NpuN, NpuC) (Gramespacher et al., 2017). The cages are connected to the split intein via cathepsin B-responsive linkers, preventing fragment association and consequent protein trans-splicing. In cancer cells overexpressing cathepsin B, the linkers would be cleaved, thus uncaging the inteins and enabling association of NpuN and NpuC. In the subsequent protein trans-splicing reaction NpuN and NpuC cleave themselves out of the construct, linking two dCas9 proteins to each other.
With these approaches, we aim to induce structural and functional changes in protein-stapled DNA selectively in cancerous tissue.
Results
We used a fluorescence readout assay to validate cathepsin B cleavage of different peptide linkers. We successfully identified one linker that could be cleaved efficiently by cathepsin B in vivo.
To investigate cathepsin B cleavage of different linkers, we incorporated five peptide linkers from literature (GFLG, FFRG, FRRL, VA, FK) (Jin et al., 2022; Shim et al., 2022; Wang et al., 2024) in between the DNA binding domain (DBD) of Gal4 and the transactivator domain VP64 (as previously described in Muench et al. (2023)). Binding of Gal4-DBD upstream of a gene encoding the fluorescence protein mCherry induces overexpression of mCherry by VP64. Consequently, separation of Gal4-DBD and VP64 by cathepsin B cleavage of the peptide linker reduces mCherry expression (see fig. 13).
Figure 13: Schematic Illustration of the Cathepsin B Fluorescence Readout Assay.The DNA binding domain (DBD) of Gal4 is conjugated to the transactivator domain VP64 via a cathepsin B-cleavable peptide linker. Binding of the Gal4-DBD to the upstream activating sequence (UAS) in proximity to the mCherry gene induces mCherry overexpression via VP64. Cathepsin B cleavage of the linker separates Gal4-DBD and VP64 and consequently reduces mCherry expression.
We conducted fluorescence readout assays in HEK293T cells. The transfected plasmids encoded mCherry, the Gal4-VP64 constructs with different linkers, and cathepsin B (see fig. 14). Additionally, a stuffer plasmid and a plasmid encoding eGFP were transfected for normalization. By conducting preliminary tests we determined adding doxorubicin 24 hours after transfection in a final concentration of 500 nM to the cell supernatant as the optimal procedure. This induces the lysosomal escape of mature cathepsin B (Bien et al., 2004). 48 hours after transfection, we measured the fluorescence intensities of mCherry and eGFP and took micrographs of the transfected cells with a fluorescence microscope.
Figure 14: Transfection Plan of HEK293T Cells for Fluorescence Readout Experiments. HEK293T cells in a 96-well plate were transfected with plasmids encoding mCherry, the Gal4-VP64 constructs with different linkers, and cathepsin B (CatB). Additionally, a stuffer plasmid and a plasmid encoding eGFP were transfected for normalization.
Figure 15 shows the fluorescence intensity of mCherry for five different peptide linkers (GFLG, FFRG, FRRL, VA, FK). The negative control was not transfected with the plasmid encoding cathepsin B. We investigated two different test conditions, in which we either transfected 30 ng or 60 ng of the plasmid encoding cathepsin B. The fluorescence intensity of mCherry was normalized by the measured fluorescence intensity of eGFP in each condition. Additionally, the values for 30 ng and 60 ng cathepsin B were normalized against the corresponding negative controls. One data point for the VA linker, transfected with 60 ng of the plasmid encoding cathepsin B, was excluded due to severe deviation from the other values. We conducted a two-way analysis of variance (ANOVA) to assess the significance of the observed differences between the negative control and the test conditions for each linker. As the negative control did not contain the plasmid encoding cathepsin B, we expected the measured fluorescence intensity of mCherry to be the highest in these conditions. However, this was only observed for the GFLG and FK linkers. Contrary to our expectations, the fluorescence intensity of the negative control was the lowest out of the three conditions tested for the remaining linkers. It appears that the addition of the plasmid encoding cathepsin B increases mCherry fluorescence intensity when the linker is not cleaved. However, this increase is only significant for the FFRG linker in the 60 ng condition. For the GFLG linker, we observed significant decreases in fluorescence intensity between the negative control and both test conditions, with no difference between the 30 ng and 60 ng conditions. For the FK linker, no significant decreases in fluorescence intensity between the negative control and the test conditions were observed.
Figure 15: Fluorescence Readout After 48 Hours for Five Different Peptide Linkers and Three Different Conditions. The fluorescence intensity for mCherry was measured for five different linkers and normalized against a baseline eGFP fluorescence intensity. The negative control was not transfected with the plasmid encoding cathepsin B. The fluorescence intensity of the negative control was set to one. Two different test conditions were investigated, in which either 30 ng or 60 ng of the plasmid encoding cathepsin B were transfected. The fluorescent readout was analyzed using a two-way ANOVA. Medium: DMEM (10% FCS). P values: ns, P > 0.05; *, P ≤ 0.05; **, P ≤ 0.01; ***, P ≤ 0.001; ****, P ≤ 0.0001.
Figure 16 shows micrographs taken with a fluorescence microscope of three different conditions: the null control, the negative control and the test sample. Figure 17 shows the corresponding graphs. All samples were transfected with plasmids encoding eGFP and mCherry. The null control and the negative control were not transfected with the plasmid encoding cathepsin B. The null control was also not transfected with any of the plasmids encoding Gal4-Linker-VP64 constructs. The test sample was transfected with 30 ng of the plasmid encoding cathepsin B and with the plasmid encoding Gal4-GFLG-VP64. As expected, the null control exhibited no detectable mCherry signal, with corresponding fluorescence intensity measurements at baseline levels. Since no plasmid encoding a Gal4-V64 construct was transfected, mCherry overexpression via VP64 could not be induced. However, we observed a high fluorescence intensity for eGFP, indicating that the transfection was successful. The negative control showed strong signals of both mCherry and eGFP. Therefore, it can be assumed that the transfection was successful and that our mCherry readout system is functional. Interestingly, there are some cells which either seem to only express mCherry or eGFP and some cells that show no fluorescence signal. The test sample showed less eGFP and mCherry fluorescence compared to the negative control. We expected to observe reduced fluorescence intensity of mCherry, as the transfected cells would express cathepsin B, which cleaves the linker, thereby decreasing mCherry expression.
Figure 16: Micrographs of HEK293T Cells in Two Control Conditions and One Test Condition. Micrographs were taken with a fluorescence microscope 48 hours after transfection. An overlay of brightfield, eGFP and mCherry is shown. All samples were transfected with plasmids encoding eGFP. The null control and the negative control were not transfected with the plasmid encoding cathepsin B. The null control was also not transfected with any of the plasmids encoding Gal4-Linker-VP64 constructs. The test sample was transfected with 30 ng of the plasmid encoding cathepsin B and with the plasmid encoding Gal4-GFLG-VP64. The micrograph of the test sample is not from the same biological replicate as the micrographs of the two controls.
Figure 17: Fluorescence Readout After 48 Hours for Two Control Conditions and One Test Condition. The fluorescence intensity for mCherry was measured for the GFLG linker and normalized against a baseline eGFP fluorescence intensity. The null control and the negative control were not transfected with the plasmid encoding cathepsin B. The null control was also not transfected with any of the plasmids encoding Gal4-Linker-VP64 constructs. The test sample was transfected with 30 ng of the plasmid encoding cathepsin B and with the plasmid encoding Gal4-GFLG-VP64.
Figure 18 shows a western blot of the wild-type (wt) version of cathepsin B as well as the truncated and mutated version of cathepsin B (Δ1-20, D22A, H110A, R116A). The truncated and mutated version of cathepsin B lacked an N-terminal signal peptide responsible for co-translational targeting to the rough endoplasmic reticulum. This would lead to the cytoplasmic expression of cathepsin B (Müntener et al., 2005). Additionally, three point mutations would disrupt the conformation of an occluding loop increasing cathepsin B activity in the cytoplasm (Nägler et al., 1997). Cells of both cathepsin B versions were treated with 500 nM doxorubicin (dox) 24 hours post-transfection and incubated for additional 24 hours. For each condition, three replicates were blotted. We observed no differences in protein expression levels between the dox-treated and untreated wt versions of cathepsin B. For the truncated and mutated version of cathepsin B, however, only the untreated samples showed the corresponding band at approximately 36 kDa expected for this version of cathepsin B. Additionally, the bands of the truncated and mutated version appeared much weaker than the ones of the wt, indicating poorer protein expression. The household protein β-tubulin is visible in all samples at approximately 55 kDa. The wt cathepsin B additionally showed bands for pro-cathepsin B at approximately 42 kDa, a mature single-chain version of cathepsin B at approximately 33 kDa and a mature double-chain version at approximately 26 kDa.
Figure 18: Western Blot of Two Versions of Cathepsin B With and Without Doxorubicin. From left to right: protein ladder, wild-type (wt) cathepsin B with (+) and without (-) doxorubicin, truncated and mutated version of cathepsin B with (+) and without (-) doxorubicin. The household protein, β-tubulin, is visible in all samples at 55 kDa. The wt cathepsin B also shows bands for pro-cathepsin B at 42 kDa, mature single-chain cathepsin B at 33 kDa and mature double-chain cathepsin B at 26 kDa. The band for the truncated and mutated version of cathepsin B can be seen in the samples without doxorubicin at 36 kDa.
Discussion
We validated the cleavage of peptide linkers by cathepsin B using a fluorescence readout assay. This assay employed a Gal4-VP64 system, where cleavage of the peptide linker by cathepsin B reduced mCherry expression, providing a reliable measure of cleavage efficiency. Five different peptide linkers (GFLG, FFRG, FRRL, VA, FK) were tested. A two-way ANOVA revealed a significant reduction in fluorescence between the negative control and the two test conditions for the GFLG linker in the presence of cathepsin B, demonstrating efficient cleavage. A significant increase in fluorescence was observed between the negative control and 60 ng test condition for the FFRG linker. However, since the increase between the negative control and the 30 ng test condition of the same linker was not significant, this difference is likely due to biological variability between the samples.
Fluorescence microscopy further supported these findings, with reduced mCherry fluorescence intensity observed in cells overexpressing cathepsin B. It was also observed that not all cells were fluorescent in accordance with an expected transfection efficiency. We also noticed that some cells seemed to only express eGFP or mCherry. However, most cells seemed to express both eGFP and mCherry as indicated by their yellow fluorescence.
Furthermore, western blot analysis confirmed that overexpressed wild-type cathepsin B was processed into its mature single-chain and double-chain forms as previously reported in the literature (Mentlein, Hattermann, Held-Feindt, 2012). This confirms that active cathepsin B is present inside of the cells. However, since lysis of the cells also disrupts the lysosomes, we can not conclude whether this active cathepsin B is also present in the cytosol in vivo. For the truncated and mutated version of cathepsin B, we only observed protein bands in samples that were not treated with doxorubicin. The three samples incubated with doxorubicin for 24 hours showed only faint bands for the housekeeping protein β-tubulin, indicating generally low protein levels. This suggests that the cells were subjected to stress, possibly through doxorubicin or inadequate handling. Along with low transfection efficiency, these factors may have contributed to the low protein levels observed.
In conclusion, these findings demonstrate that our fluorescence-based readout assay can reliably detect cathepsin B-mediated cleavage of peptide linkers, with the GFLG linker showing particular susceptibility to cleavage. This makes GFLG a promising candidate for targeted applications in environments with upregulated cathepsin B activity, such as in cancerous tissues. Additionally, our cathepsin B-cleavable linker can be combined with caged inteins (Gramespacher et al., 2017) conjugated to a dead Cas9 to selectively induce Cas-stapling in the presence of cathepsin B.
Outlook
Future experiments could investigate the influence of different doxorubicin concentrations on the activity of cathepsin B in the cytosol. Different linker lengths or a repeat of the GFLG linker could also be tested. Additionally, this system could be used for other proteases that are involved in certain diseases, such as different caspases in neurodegenerative conditions (Espinosa-Oliva et al., 2019).
Readout Systems
By developing the EMSA and FRET assays, we established key tools for the rational design and evaluation of various protein staples. Beginning with the successful construction of the basic staples, these assays provided key insight into DNA binding proteins and stapling mechanisms. Through thorough testing of our workflow, we developed foundational techniques that future iGEMers and researchers can leverage to engineer and optimize protein-based DNA-folding systems.
Introduction
Selection of DNA binding Proteins tetR, Oct1 and GCN4
Both assays were initially established in a simple system using the Tetracycline Repressor (tetR)
and the
human transcription factor Oct1 as they are well-characterized proteins with known binding
properties.
TetR is a bacterial transcriptional repressor that binds specifically to the tetO operator
sequence and
dissociates in the presence of tetracycline. It is widely adopted as a synthetic gene regulation
tool,
both in eukaryotic and prokaryotic systems (Berens & Hillen, 2004). In a similar manner, Oct1, a Pit-Oct-Unc
(POU)
domain transcription factor involved in immune cell regulation and stress response, has been shown
to bind
tightly to its octamer DNA motif (Lundbäck et al., 2000; Stepchenko
et al., 2021). It was shown that the DNA binding domain of Oct1 can be
readily fused
to other proteins for increased protein
solubility and strong DNA binding capabilities, even during protein purification (J. H. Park
et al., 2013; Y. Park et al., 2020). We genetically
fused tetR
and Oct1 to create
the "simple staple".
In addition to tetR and Oct1, we also looked into small basic-region leucine zipper (bZip)
proteins with
DNA binding capabilities. The motif consists of a coiled-coil leucine zipper dimerization domain,
and a
highly charged basic region that directly contacts and binds to DNA (Hollenbeck & Oakley, 2000).
One well
characterized example is the General Control Protein 4 (GCN4), a well-characterized
transcriptional
activator from yeast (Arndt & Fink, 1986). At its N-terminus, GCN4 contains basic residues, the
so-called
bZip domain, through which it binds specifically to the cyclic AMP response element (CRE) DNA
sequence
(Hollenbeck et al., 2002). A variant of GCN4 with the DNA binding bZip-domain at the
C-terminus
(rGCN4) has been engineered to bind to the inverted CRE sequence, INV2 with similar affinity
(Hollenbeck
et al., 2001). By genetically fusing GCN4 to rGCN4, we created a small bivalent DNA binding
staple with less than 150 amino acids.
Förster Resonance Energy Transfer (FRET)
Förster Resonance Energy Transfer (FRET) is a distance-dependent physical process where energy is
transferred non-radiatively from an excited donor fluorophore to an acceptor fluorophore via
dipole-dipole
coupling. The efficiency of energy transfer is highly sensitive to the distance between the donor
and
acceptor, typically in the range of 1-10 nm, making FRET ideal for studying molecular proximity
(Hochreiter et al., 2019). This proximity sensitivity is governed by the Förster radius
(R0),
which is the
distance at which 50 % energy transfer occurs. Factors affecting FRET efficiency include the
overlap of
the
donor's emission spectrum with the acceptor's absorption spectrum, the quantum yield of the donor,
and the
relative orientation of the fluorophores (Wu & Brand, 1994). These characteristics allow to
detect
interactions such as protein-DNA binding or DNA proximity in real time with FRET.
For our assay, we selected mNeonGreen and mScarlet-I as donor and acceptor, respectively, due to
their
strong fluorescence, spectral overlap, and minimal photobleaching, ensuring high FRET efficiency
in our
system (Bindels et al., 2017; Shaner et al., 2013).
FRET's
sensitivity to small changes in distance makes
it especially powerful for analyzing molecular interactions in living cells (Okamoto & Sako,
2017).
Electrophoretic Mobility Shift Assay (EMSA)
The Electrophoretic mobility shift assay (EMSA) is a widely adopted method used to study DNA-protein interactions. EMSa functions on the basis that nucleic acids bound to proteins have reduced electrophoretic mobility, compared to their counterpart. (Hellman & Fried, 2007). Mobility-shift assays can both be used to qualitatively assess DNA binding capabilities or quantitatively to determine binding stoichiometry and kinetics such as the apparent dissociation constant (Kd) (Fried, 1989).
Aim of this subproject
Engineering a solid and versatile toolbox is a huge challenge, especially when working with complex Cas staples. To systematically characterize these systems, we first set out to develop a well-tested collection of assays that lay the foundation to characterize the more complex aspects of our PICasSO toolbox. These assays provide essential tools for studying protein-DNA interactions and proximity. We used electrophoretic mobility shift assays (EMSA) to analyze binding kinetics and sequence specificity in vitro and a Förster resonance energy transfer (FRET) assay to detect DNA-DNA proximity in vivo. Together, these assays form the backbone of our experimental approach and were utilized to systematically analyze and create new staples.
Results
The FRET assay was developed using a two-plasmid system in bacterial cells.After testing different constructs, our final expression plasmid
contains a tetR binding site and expresses three key proteins under the control of a single T7
promoter in a polycistronic operon: (1) tetR-Oct1, our simple staple fusion protein that acts as a
bivalent DNA binding protein, tethering two plasmids via tetR and Oct1 binding sites; (2)
Oct1-mNeonGreen, serving as the FRET donor; and (3) tetR-mScarlet-I, the FRET acceptor. This
ensures all three proteins are co-expressed in similar stoichiometry. The folding plasmid contains
an Oct1 binding site for the staple and FRET donor binding.
When tetR-Oct1 binds the respective sites on both plasmids, mNeonGreen and mScarlet-I are brought into
into proximity, facilitating FRET between the two fluorophores. Successful stapling of the
plasmids results in increased energy transfer from mNeonGreen to mScarlet-I, which can be detected
by exciting mNeonGreen and measuring enhanced emission from mScarlet-I. A positive control,
consisting of a direct fusion of mNeonGreen and mScarlet-I, ensures maximal FRET efficiency and
serves as a benchmark for the assay.
Fluorescence intensity, normalized to cell count, of mNeonGreen and mScarlet-I was measured 18 h
after induction with varying IPTG concentration (see fig. 19A and 19B). An increasing
expression rate is visible for decreasing IPTG concentrations. Fluorescence intensity of the positive control was
significantly stronger compared to the negative control and staple. The negative control and
staple, which both have the same expression plasmid construct, had similar fluorescence intensity
for mNeonGreen and mScarlet-I down to approximately 0.05 mM. Lower concentrations resulted in
strong discrepancies. To ensure comparability between the negative control and staple, further
fluorescence intensity measurements were conducted after induction with 0.05 mM IPTG. Fluorescence
measurement of the donor and acceptor showed similar intensities, with only a small significant
difference for mNeonGreen. A large difference could be measured between the staple and negative
control, indicating proximity induced FRET (see fig. 19C).
Figure 19: Fluorescence measurement of mNeonGreen, mScarlet-I and FRET. Fluorescent measurement, normalized to cell count, of mNeonGreen (ex. 490 nm, em. 530 nm), mScarlet-I (ex. 560 nm, em. 600 nm), and FRET (ex. 490 nm, em. 600 nm) in E. coli, 18 h after induction with 0.025 mM IPTG. Data is presented as mean +/- SD. A, B Fluorescence intensity of mNeonGreen and mScarlet-I with different IPTG concentrations. C Fluorescence intensity of FRET pair after induction with 0.05 mM IPTG. (n = 3) Statistical significance was determined with Ordinary two-way ANOVA with Šidák's multiple comparison test, with a single pooled variance. *p < 0.05, ****p < 0.001. Only significant results are shown.
The DNA binding proteins tetR and Oct1-DBD were fused to mScarlet-I and mNeonGreen, respectively, and to each other, each harboring a His6-tag. The bZip proteins GCN4, rGCN4 and their fusion bGCN4 were fused N-terminally to a FLAG-tag (DYKDDDDK). All proteins could be readily expressed under the T7 promoter in E. coli BL21 DE3 and purified with Ni-NTA, or Anti-FLAG affinity columns for His-tagged and FLAG-tagged proteins, respectively (see figure 20).
Figure 20: SDS-PAGE analysis of purified DNA binding proteins. A) Analysis of fractions eluate of purified protein taken during Ni-NTA affinity chromatography. b) Analysis of fractions eluate of purified protein taken during Anti-FLAG affinity chromatography 1 µL of each sample was prepared with Leammli buffer and loaded on 4-15% TGX-Gel. Correct bands of interest are highlighted by red boxes
To first assess possible DNA binding and test out different buffer systems qualitative EMSA was for the purified proteins and additionally three different buffer systems tested for Oct1-DBD and tetR. DNA binding could be detected for the single purified proteins, but not for the bGCN4 fusion (see fig. 21). Binding buffer 1 (137 mM NaCl, 2.7 mM KCl, 10 mM Na 2HPO4, 1.8 mM KH2HPO4, 0.1 % (v/v) IGEPAL® CA-360, 1 mM EDTA), also described by Hollenbeck (2001) was the best performing buffer and used for subsequent experiments.
Figure 21 Qualitative EMSA results Electrophoresis was performed in TBE buffer with 10 % TGX-Gel pre-equilibrated with TBE, bands are visualised by post-staining with SYBR-Safe A) Purified mNeonGreen-Oct1 (1000 nM, 100 nM or 10 nM) or tetR-mScarlet-I (1000 nM) were equilibrated, in different buffer compositions, with 0.5 µM DNA containing three Oct1 or tetR binding sites respectively. (Binding buffer 1: 137 mM NaCl, 2.7 mM KCl, 10 mM Na 2HPO4, 1.8 mM KH2HPO4, 0.1 % (v/v) IGEPAL® CA-360, 1 mM EDTA; Binding buffer 2: 10 mM Tris, 50 mM KCl; NaP250: 50 mM NaH2PO4, 150 mM NaCl, 250 mM Imidazol) B) Purified tetR-Oct-1 fusion protein was incubated with 0.5 µM DNA containing either a tetR or Oct-1 binding site C) 200 µM purified protein were equilibrated with 0.5 µM DNA containing one target site. Bands were visualized with SYBR-Safe staining.
To further analyze DNA binding, quantitative shift assays were performed for GCN4 and rGCN4. Here
0.5 µM DNA were incubated with varying concentrations of protein until equilibration. After
electrophoresis, bands were stained with SYBR-Safe and quantified based on pixel intensity. The
obtained values were fitted to equation 1, describing formation of a 2:1 protein-DNA complex:
Here [L]tot describes the total protein monomer concentration, Ka corresponds to the apparent monomeric equilibration constant. The Θmin/max values are the experimentally determined site saturation values (For this experiment 0 and 1 were chosen for min and max respectively). GCN4 binds to its optimal DNA binding motif with an apparent dissociation constant Kk of (0.2930.033)×10-6 M, which is almost identical to the rGCN4 binding affinity to INVii a d of (0.2980.030)×10-6 M (see fig. 22).
Figure 22: Kd Calculation of GCN4 and rGCN4 Quantitative assessment of binding affinity for GCN4 and rGCN4. Proteins of varying concentrations were incubated with 0.5 µM DNA in Binding buffer 1, and the bound fraction analyzed by dividing pixel intensity of bound fraction with pixel intensity of bound and unbound fraction using ImageJ. At least three separate measurements were conducted for each data point. Values are presented as mean +/- SD
Discussion
The results from the fluorescence intensity measurements showed stronger expression of the fluorescent
proteins for the positive control. Based on our preliminary testing, explained in our engineering
cycles(#link#), we suspect a strong metabolic burden to influence expression levels, especially for the
staple construct. This could be both due to the polycistronic expression of multiple protein, as well as
the strong T7 promoter resulting in too high mRNA levels. The negative control and
staple showed
strong discrepancies in mNeonGreen and mScarlet-I fluorescence for IPTG concentrations below 0.05 mM. This
is surprising as both samples have the same expression plasmid and only the folding plasmid of the
negative control is missing the binding site for Oct1. One possible explanation could be off-target
binding in the E. coli genome, resulting in changes in the expression pattern or
similar. With 84 specific Oct1 binding sequences previously reported in the genome this could be a major
factor. (Y. Park et al., 2020). Studies also showed binding affinity of Oct1 to
different target sites,
albeit with lower affinity, possibly resulting in even more potential genomic binding sites (Verrijzer et al., 1992). The folding plasmid, harboring a p15A origin of replication,
is expected to have around eleven copies per cell (Shao et al., 2021), resulting
in approximately 130 Oct1 binding sites, which stand in competition to genomic binding sites.
Further experiments are needed to better understand this observation.
All proteins for in vitro characterization could be expressed and purified. Some nonspecific proteins
still remained in the eluate, but this is to be expected given well known unspecific binding of cellular
proteins to the Ni-NTA affinity matrix. Initial qualitative tests showed successful binding of the single
DNA binding proteins and the tetR-Oct1 fusion to the DNA. Since tetR and Oct1 were fused with
mScarlet-I and
mNeongreen, respectively, we could show that these proteins accept fusions to proteins and can still bind
to DNA, which also matches previous results shown in literature (Gossen & Bujard, 1992; J. H. Park et al.,
2013b). Bound and unbound fractions were visualized on the gel by SYBR-Safe staining and illumination with
a trans-illuminator, resulting. To ensure good quantifiability, and to further characterize additional DNA
binding proteins the bZip proteins were chosen for quantitative gel shift assays. For the purified
proteins GCN4, rGCN4 and the fusion bGCN4 a qualitative gel was run with high protein concentration. Bands for GCN4 and rGCN4 were visible but no band for bGCN4 could be
detected, indicating a
lack of DNA binding. This suggests that the dimerization, necessary for DNA binding, is disrupted by the
GSG-linker (Ellenberger et al., 1992; Liu et al., 2006; Lupas
et al., 2017; Woolfson, 2023). To better
understand possible problems in dimerization circular dichroism can be used to analyze secondary structure
and proper coiled coil formation (Greenfield, 2006). Further engineering can be done by testing out
various linkers with specific properties to ensure correct folding and dimerization (Chen et al., 2013).
The apparent binding kinetics calculated for GCN4 ((0.2930.033) × 10-6 M) and rGCN4
((0.2980.030) × 10-6 M) are
approximately a factor 10 higher then those described in literature ((96) × 10-8 M for
GCN4 and (2.90.8) × 10-8 M for rGCN4) (Hollenbeck et al., 2001). The
differences could be due to the lower sensitivity of SYBR-Safe
staining compared to radio-labeled oligos.
The FLAG-tag fusion to the N-terminus of proteins could potentially decrease binding affinity, likely due
to steric hindrance affecting the interaction with DNA. Interestingly, the differences in binding affinity
between GCN4 and rGCN4 appear negligible. Since GCN4 binds to DNA via its N-terminus and rGCN4 binds
C-terminally, the FLAG-tag likely does not directly influence DNA binding. However, it may influence the
dimerization of the proteins, which is necessary for DNA binding. To further investigate this, the
FLAG-tag can be cleaved using an enterokinase and potential changes in binding affinity analyzed.
Outlook MMMM
MARIK WILL SAVE THE WORLD.
Delivery System
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Introduction
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The CRISPR / Cas System as a Gene Editing Tool
It has long been known that the three-dimensional organization of DNA has major implications in many contexts from plasmid delivery into cells over the transcriptional activity of genes to the spatial organization of chromosomes or whole genomes. Importantly, structural and orientational rearrangements of DNA are associated with several genetic diseases including cardiomyopathy and cancer (Dong et al., 2023) (Watanabe et al., 2020). Being able to efficiently engineer the spatial genome organization or even defined DNA structures in vivo would revolutionize our ability to understand and control cellular systems and address diseases related to chromatin organization defects.
The CRISPR / Cas System as a Gene Editing Tool
It has long been known that the three-dimensional organization of DNA has major implications in many contexts from plasmid delivery into cells over the transcriptional activity of genes to the spatial organization of chromosomes or whole genomes. Importantly, structural and orientational rearrangements of DNA are associated with several genetic diseases including cardiomyopathy and cancer (Dong et al., 2023) (Watanabe et al., 2020). Being able to efficiently engineer the spatial genome organization or even defined DNA structures in vivo would revolutionize our ability to understand and control cellular systems and address diseases related to chromatin organization defects.
The CRISPR / Cas System as a Gene Editing Tool
It has long been known that the three-dimensional organization of DNA has major implications in many contexts from plasmid delivery into cells over the transcriptional activity of genes to the spatial organization of chromosomes or whole genomes. Importantly, structural and orientational rearrangements of DNA are associated with several genetic diseases including cardiomyopathy and cancer (Dong et al., 2023) (Watanabe et al., 2020). Being able to efficiently engineer the spatial genome organization or even defined DNA structures in vivo would revolutionize our ability to understand and control cellular systems and address diseases related to chromatin organization defects.
Results
Lorem ipsum dolor sit amet consectetur adipisicing elit. Corrupti doloremque necessitatibus, praesentium fuga labore ipsum dolores architecto laudantium voluptatum dolorem eius ducimus porro velit expedita non sint illo sequi rem.
Lorem ipsum dolor sit amet consectetur adipisicing elit. Corrupti doloremque necessitatibus, praesentium fuga labore ipsum dolores architecto laudantium voluptatum dolorem eius ducimus porro velit expedita non sint illo sequi rem.
Lorem ipsum dolor sit amet consectetur adipisicing elit. Corrupti doloremque necessitatibus, praesentium fuga labore ipsum dolores architecto laudantium voluptatum dolorem eius ducimus porro velit expedita non sint illo sequi rem.
Lorem ipsum dolor sit amet consectetur adipisicing elit. Corrupti doloremque necessitatibus, praesentium fuga labore ipsum dolores architecto laudantium voluptatum dolorem eius ducimus porro velit expedita non sint illo sequi rem.
Lorem ipsum dolor sit amet consectetur adipisicing elit. Corrupti doloremque necessitatibus, praesentium fuga labore ipsum dolores architecto laudantium voluptatum dolorem eius ducimus porro velit expedita non sint illo sequi rem.
Discussion
Editing Efficiency of fgRNAs Is Influenced by Numerous Factors
It has long been known that the three-dimensional organization of DNA has major implications in many contexts from plasmid delivery into cells over the transcriptional activity of genes to the spatial organization of chromosomes or whole genomes. Importantly, structural and orientational rearrangements of DNA are associated with several genetic diseases including cardiomyopathy and cancer (Dong et al., 2023) (Watanabe et al., 2020). Being able to efficiently engineer the spatial genome organization or even defined DNA structures in vivo would revolutionize our ability to understand and control cellular systems and address diseases related to chromatin organization defects.
Fusion Guide RNA based Cas Staples Facilitate a Promising New Approach of 3D Genome Engineering
It has long been known that the three-dimensional organization of DNA has major implications in many contexts from plasmid delivery into cells over the transcriptional activity of genes to the spatial organization of chromosomes or whole genomes. Importantly, structural and orientational rearrangements of DNA are associated with several genetic diseases including cardiomyopathy and cancer (Dong et al., 2023) (Watanabe et al., 2020). Being able to efficiently engineer the spatial genome organization or even defined DNA structures in vivo would revolutionize our ability to understand and control cellular systems and address diseases related to chromatin organization defects.
Outlook
It has long been known that the three-dimensional organization of DNA has major implications in many contexts from plasmid delivery into cells over the transcriptional activity of genes to the spatial organization of chromosomes or whole genomes. Importantly, structural and orientational rearrangements of DNA are associated with several genetic diseases including cardiomyopathy and cancer (Dong et al., 2023) (Watanabe et al., 2020). Being able to efficiently engineer the spatial genome organization or even defined DNA structures in vivo would revolutionize our ability to understand and control cellular systems and address diseases related to chromatin organization defects.
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