Self-inflicted DNA damage drives cancer cell resistance to radiation therapy
Commonly used radiation therapy kills cancer cells by inflicting extensive DNA damage to the irradiated tissue. Naturally occurring DNA damage in human cells is efficiently remedied by a large number of DNA repair pathways. Yet this process takes time, and the addiction of cancer cells to rapid uncontrolled cell division renders them incapable of dealing with the level of DNA damage inflicted by high doses of radiation. As a result, radiation therapy efficiently kills tumor cells. Despite the success of the method, reoccurrence of tumors is still common. The mechanisms through which tumor cells manage to evade cell death after lethal doses of irradiation are not well understood, and hence resistance to RT remains a considerable challenge for effective clinical tumor control.
An international team including researcher from our group at the Department of Medical Biochemistry and Biophysics, Division of Genome Biology, Karolinska Institutet now shed light into an unexpected strategy of cancer cells to evade death by radiation in a study published in Science. The team of scientists working in Denmark, Sweden, Canada and Switzerland, coordinated by Claus Storgaard Sørensen at the Biotech Research & Innovation Centre (BRIC), University of Copenhagen, found that in response to radiation, tumor cells can activate an endogenous nuclease, CAD, which initiates genome-wide DNA breaks. While persistent DNA damage is generally bad news for the cell, the self-inflicted DNA breaks made in this context helps cancer cells to suspend their program to divide, pausing their cell cycle at the so-called G2 checkpoint, and gain time to repair the remaining DNA damage.
Using a method to map the ‘nicks’ – single-strand DNA breaks – introduced by activation of CAD in response to radiation, researchers at SciLifeLab then found that self-inflicted DNA damage was non-random and instead concentrated on a small number of regions in the genome. What’s more, the authors found that this phenomenon was specific for cancer cells as loss of CAD activity rendered cancer cells, but not normal cells, vulnerable to radiation-induced damage.
“With CAD-inflicted DNA damage following a discernable pattern, we hope to investigate in the future how the cell targets and contains this potent endogenous nuclease activity.” says Philip Yuk Kwong Yung, postdoctoral fellow in our lab.
Collectively, the findings highlight a cancer-specific survival mechanism that could be targeted and exploited to enhance the tumor cells’ vulnerability to genotoxic cancer treatments. Indeed, the study also showed that experimental blocking of the CAD function made tumor cells (but not normal healthy cells) more sensitive to radiation, thereby suggesting how this new knowledge could be used to improve the outcome of radiotherapy in the future.
Our research was funded by Vetenskapsrådet and ERC.
From supplementary information of recent JACS paper
Figure 1 – Labeling of SARS-CoV-2 derived microproteins in fixed cells.
COS-7PylRS-AF cells were transfected with the indicated constructs and incubated for 24 h with 50 µM TCO*K. Cells were subsequently fixed, permeabilized and labeled with met-tet-BDP-FL and analyzed by confocal microscopy. SARS-CoV-2 M (Membrane) protein was used as a control because of its known localization to the Golgi (Klumperman et al., 1994). ORF7b is a type I (luminal N-terminus) transmembrane protein implicated as virulence factor, generated by leaky ribosome scanning in an alternative frame within the main ORF7a (Pfefferle et al., 2009). The homologous microprotein in SARS1 is suggested to localize to the Golgi (Schaecher et al., 2008). Our observation is a wider distribution in Golgi, ER and plasma membrane. ORF9b is an alternative ORF within ORF9a and the SARS1 homolog is reported to localize to mitochondria (Shi et al., 2014). We also observe a distinct speckled pattern in the cytosol compatible with partial mitochondrial localization. ORF9c expression and function is currently unknown but we observe a distinct localization with cellular membranes. ORF3b of SARS-CoV-2 is truncated to a 22 amino acid fragment due to a premature stop codon as compared to the SARS1 homolog, but has been reported to potently suppress host cell antiviral interferon response (Konno et al., 2020). The truncation preserves a single predicted transmembrane domain and our imaging suggests an association with cellular membranes.
Movie 1: Widefield, live cell imaging of SARS-CoV-2-ORF6N-TCO*K. 24 h after transfection and TCO*K incubation, COS-7PylRS-AF cells were exposed for 30 min to 500 nM me-tet-BDP-FL. Right after removal of the dye, cells were imaged over a short period of time. In the movie, it is possible to see ORF6-containing vesicles travelling through the cytoplasm. Timestamps (min:sec) are displayed in the top-right corner of each frame.
RESOURCES
ID
Name
Reference/Repository
E407
pAS1_4xh7SKPylT_EF1_PylRS AF_IRES_Puro
Addgene 140023
K122.5
pAS1_4x7SKPylT_EF1_Ub-*CoV2_M
Addgene 162815
K124.1
pAS1_4x7SKPylT_EF1_Ub-*CoV2_9b
Addgene 162816
K125.2
pAS1_4x7SKPylT_EF1_Ub-*CoV2_ORF6
Addgene 162817
K126.1
pAS1_4x7SKPylT_EF1_Ub-*CoV2_ORF7b
Addgene 162818
K127.2
pAS1_4x7SKPylT_EF1_Ub-*CoV2_ORF10
Addgene 162819
K128.2
pAS1_4x7SKPylT_EF1_Ub-*CoV2_ORF9c/14
Addgene 162820
K132.2
pAS1_4x7SKPylT_EF1_Ub-*CoV2_ORF3b_alt
Addgene 162821
K134.10
pAS1_4x7SKPylT_EF1_Ub-*CoV2_ORF3b(CoV1h)
Addgene 162822
Table 1 – Plasmids are available on request and via Addgene:
METHODS
COS-7 cell lines were maintained in Dulbecco’s Modified Eagle Medium (DMEM), containing high glucose, GlutaMAXTM and pyruvate (Gibco). Medium was supplemented with 10% fetal bovine serum (FBS, Sigma-Aldrich). All cell lines were cultured in an ambient-controlled incubator at 37°C, 5% O2 and 5% CO2. Cells were forward transfected using Lipofectamine LTXTM with PLUSTM reagent (Invitrogen) according to manufacturer’s protocol. Axial trans-cyclooct-2-ene-l-lysine (TCO*K) was added at the time of transfection as indicated and cells were harvested after 24 h. TCO*K (SiChem, SC-8008) stock solution was prepared at 100 mM in 0.2 M NaOH/H2O, 15% DMSO. 6-Methyl-Tetrazine-BODIPY®-FL (me-tet-BDP-FL, Jena Bioscience) stocks were prepared in DMF and further diluted in either RIPA buffer (lysate labeling), TBS-T (fixed cells labeling) or the appropriate growth medium (live cell labeling).
Immunofluorescence and live cell imaging
For immunofluorescence, cells were grown and transfected in 96-well µ-Plates (ibidi). After withdrawing the ncAA for 4 h, cells were fixed in 4% formaldehyde for 10 min at room temperature and permeabilized for 15 min with 0.1% (v/v) triton/PBS. Prior to incubation with the appropriate antibodies, cells were click-labeled with 500 nM 6-Methyl-tetrazine-BODIPY-FL (Jena Bioscience), washed 3 times with PBS and then blocked for 1 hour in 2% BSA in TBS supplemented with 0.1% Tween-20 (TBS-T). Cells were incubated with the HA-probe antibody (F-7, Santa Cruz Biotechnology) overnight at 4°C. After washing with TBS-T, cells were stained with Alexa555-conjugated secondary antibodies (Life Technologies) for 60 min at room temperature and counterstained with 1 mg/ml DAPI (Sigma-Aldrich). After washing, cells were imaged on a Zeiss LSM780 confocal laser scanning microscope using a 40x/1.3 oil objective.
For live cell imaging, cells were grown and transfected in 96-well imaging plates (BD Falcon). After withdrawing ncAA for 1 hour, cells were incubated 30 minutes at 37°C in presence of 500 nM me-tet-BDP-FL. Where stated, cells were co-stained with either 4 µM Hoechst, 10 mM ER-trackerTM Red or 250 nM MitoTrackerTM Orange CMTMRos (Invitrogen). After washing 2 times with PBS, cells were immediately imaged in Live Cell imaging Solution (Molecular Probes), on a Nikon eclipse Ti2 inverted widefield microscope equipped with a heated imaging chamber. Images were acquired using a 20×/0.75 air objective or a 40x/1.15 water objective. For the long-term imaging experiment shown in Supplementary Figure 3, cells were maintained in Leibovitz’s L-15 Medium (Gibco) and imaged using a 10x/0.45 air objective.
Proteins are the molecular machines of life, performing a myriad of functions inside every cell of our body. Proteins are assembled from small building blocks, the amino acids, by large protein factories called ribosomes. Understanding how proteins work is a quest of basic biology and medical research.
Postdoctoral researcher Lorenzo Lafranchi and PhD student Dörte Schlesinger. Not in the picture is Kyle Kimler, former master student, now at Broad Institute.
To study proteins inside of human cells, researchers have been using light microscopes for more than one hundred years. Thirty years ago, the cloning of the green fluorescent protein GFP, together with genetic engineering tools, revolutionized the field by enabling researchers to fuse a fluorescent ‘beacon’ to any protein of interest so that it can be directly observed in living cells using fluorescence microscopy. Fast forward, today’s microscopes achieve live imaging, at nanometer resolution, in multicolor, allowing researchers to resolve even the smallest subcellular structures and essentially watch protein at work.
Fluorescent proteins and other tools that are available to researchers have however one limitation: the size of the fluorescent tag is often equivalent to the size of a typical folded protein, thus adding a considerable molecular ‘cargo’ to the protein under study and potentially impacting its function. This can become a particular obstacle for the study of microproteins, a newly appreciated class of proteins that are much smaller than average. Such tiny proteins have often been overlooked in the past but seminal discoveries of microproteins with important biological functions have sparked growing interest by the research community.
We have made it a focus of our laboratory to tackle the challenges of discovering and characterizing microproteins. Here, we developed a method which allows fluorescent tagging of proteins with the smallest imaginable perturbation – a single amino acid – added genetically on either end of a (micro)protein of interest.
For the method, termed STELLA, a synthetic building block (a non-canonical “designer” amino acid, rather than one of the 21 canonical ones) is incorporated together with a larger tag using a technique termed genetic code expansion. The tag however is swiftly removed by the cell, leaving a single terminal designer amino acid on the protein of interest. As an advantage over existing labeling techniques relying on genetic code expansion, STELLA can thus be used to conveniently and universally label the termini of any proteins. While very similar to its natural counterpart, the designer amino acid introduces a peculiar chemical group into the protein that subsequently allows conjugation with a small organic fluorescent dye, now lighting up the protein of interest inside of the living cell.
So-called transposons are abundant DNA-elements found in every eukaryotic organism as a consequence of their ability to jump and multiply within the host genome. Their activity represents a threat to the integrity of the host genome and thus the host cell engages a number of protective mechanisms to silence the expression of transposons. It is known that some of these mechanisms fail in cancer cells and also ageing cells, leading to a mobilization of transposons with largely unknown consequences. Histones, the proteins that package the genome in the eukaryotic nucleus, are key to the most fundamental line of defense to transposons. By forming a highly compacted array, so-called heterochromatin, they render the associated DNA sequence inert to being read and expressed. Heterochromatin is defined by characteristic modifications to histone proteins and DNA, such as histone H3 K9 trimethylation and DNA CpG methylation.
Here, we studied endogenous retroviral elements (ERVs), a particularly active and abundant family of transposable elements in the mouse genome, which are in fact remnants of once-active viruses. Curiously, while we found all the hallmarks of heterochromatin to be employed in the silencing mechanism, ERV chromatin was highly enriched in a histone variant, termed histone H3.3, which has previously been invariably associated with active regions of the genome. Following up on this observation, wecould elucidate an unexpected mechanism involving a continuous loss of ‘old’ histones and replenishment with newly synthesized histones H3.3 molecules. By genetic manipulation, we were able to deduce a mechanism explaining this dynamic process: the ATP-dependent chromatin remodeler Smarcad1 evicts histones within heterochromatin, thus creating gaps in the chromatin fibre that could render parts of the ERV gene accessible. Following suit, the histone chaperone DAXX seals these gaps by facilitating reassembly of nucleosomes with histone variant H3.3. The concerted process of eviction of one and deposition of another histone is so smooth and efficient that it leaves no apparent trace of accessible DNA.
The result is puzzling because active remodeling and nucleosome eviction is expected to counteract a compacted chromatin structure, inert to transcriptional activation. But we believe that dynamic heterochromatin is an adaption of a ubiquitous silencing mechanism to the specific requirements of a pluripotent chromatin state. The highly transient opening of heterochromatin may allow sequence-specific co-repressors to find their target DNA sequence within the transposable element, in turn recruiting more repressive factors to propagate and amplify the silent state.
Our paper in Nature Methods (“Genetic code expansion in stable cell lines enables encoded chromatin modification“, DOI:10.1038/NMETH.3701) is the first one to generate and characterize stable amber suppression cell lines for unnatural amino acid mutagenesis . The principle of genetic code expansion via amber suppression is shown below
We then apply the system to generate genetically encoded synthetic histone acetylation marks to directly test the function of this posttranslational modification in chromatin, one position at a time. This approach highlights the potential of the methodology to perform experiments with biochemical precision in living cells that could otherwise only be achieved in vitro. In vitro experiments can provide a clear link between molecular cause and effect, but are abstracted from the appreciable complexity of the cellular environment. In contrast, in vivo experiments typically provide a wealth of correlative information about changes of chromatin state in a native context, but it is commonly impossible to infer direct causation from these experiments. For example, all histone acetyl transferases (HATs) are known to acetylate a range of sites and substrates, including non-histone proteins, thus genetic knockout or enzymatic inhibition of HATs does not directly and exclusively test the function of histone acetylation. Employing a synthetic route to modulate cognate posttranslational modifications has the power to show direct causality between the modifications and their downstream effects, abstracted from the complexity of enzymes that set and erase the modifications. We believe that in the future such approaches to synthetic epigenetics will be very powerful for defining the function of posttranslational modifications, in particular the complex modification code present on histones.
In a second publication, we have employed stable amber suppression in HEK293 cell lines to synchronously activate a mutant Isocitrate-dehydrogenase enzyme (IDH2) in the entire populaiton of cells by light and followed changes in metabolic and epigenetic products:
We have received a 4-year research grant from Vetenkapsrådet for studying stem cell chromatin and a grant from Åke Wibergs Foundation for studying the role of chromatin regulators in suppressing genomic instability.
The Elsässer Lab at Karolinska Institutet 
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