Background

RNA polymerase II (RNAPII) plays an essential role in protein production by transcribing DNA into mRNA. In addition, it participates in the detection, signaling, and repair after DNA damage (Ljungman and Lane, 2004; Lagerwerf et al., 2011; Lans et al., 2019; Landsverk et al., 2021), and thus also plays a key role in the DNA damage response. Furthermore, RNAPII is essential for cell cycle progression, and is itself regulated by the cell cycle (Enserink and Chymkowitch, 2022). As DNA damage greatly affects the cell cycle by activation of cell cycle checkpoints (Hauge et al., 2021), measuring RNAPII levels on chromatin in individual cell cycle phases is highly relevant both in the absence and presence of DNA damage. Previously, cell cycle effects on RNAPII have been studied, for instance by microscopy (Chan et al., 2012; Teves et al., 2018) and/or chromatin immunoprecipitation/fractionation followed by sequencing (Liang et al., 2015, Wang et al., 2021) or western blotting (Akoulitchev and Reinberg, 1998; Palozola et al., 2017). However, although sequencing or chromatin fractionation techniques can give high resolution sequence information and/or quantitative data, they have so far been based on cell lysates made from a large number of pooled cells. In order to investigate cell cycle effects, these techniques require cell synchronization, which can induce replication stress or changes to transcription. Microscopy offers single-cell resolution and does not require synchronization but is limited in the number of cells analyzed. To overcome these obstacles, we recently described a new method to study RNAPII chromatin binding in single cells through the cell cycle (Bay et al., 2022). This flow cytometry method allows for the determination of cell cycle phases without the need to synchronize cells. Also, thousands of cells are measured per sample per experiment, ensuring accurate quantitative readouts. Using this method on cells treated with the transcriptional inhibitor 5,6-Dichloro-1-beta-Ribo-furanosyl Benzimidazole (DRB), which arrests RNAPII in promoter proximal regions, and the proteasome inhibitor MG132, we showed that promoter proximal RNAPII is degraded under unperturbed conditions, and that this is further enhanced after UV treatment (Bay et al., 2022). In line with our results, degradation of promoter proximal RNAPII after UV treatment was also observed in an independent study by live-cell imaging, chromatin immunoprecipitation followed by sequencing, and western blotting of chromatin fractions (Steurer et al., 2022). Furthermore, using our method, we could observe cell cycle differences in the chromatin loading of RNAPII after UV treatment (Bay et al., 2022). Here, we present the detailed protocol for this method.

A first critical step in the protocol is the cell extraction step, where non-chromatin-bound factors are removed by treatment with mild detergent Triton X-100 (0.5%) prior to fixation (Figure 1). This allows measurements of only chromatin-bound RNAPII. The use of flow cytometry to measure chromatin-bound factors after detergent treatment was first described in the early 90s for the study of chromatin-bound DNA polymerase α (Stokke et al., 1991). Similar assays using extraction prior to fixation have also been used to measure chromatin-binding of other proteins involved in transcription, replication, and repair by flow cytometry (Stokke et al., 1993; Forment et al., 2012; Forment and Jackson, 2015) or immunofluorescence microscopy (Syljuåsen et al., 2005; Britton et al., 2013). Notably, this is the first time such a protocol has been described for RNAPII by flow cytometry. Triton X-100 treatment is performed in a sucrose-containing buffer, similar to pre-extraction/cytoskeletal buffers commonly used in immunofluorescence microscopy (Britton et al., 2013; Kilgas et al., 2021). However, as RNAPII is known to be released from chromatin in mitosis (Parsons and Spencer, 1997), NaCl concentrations were optimized based on the expected low chromatin binding of RNAPII in mitotic cells (Bay et al., 2022). Moreover, as RNAPII is a target for proteasome-mediated degradation (Wilson et al., 2013), particular care should be taken to avoid degradation during cell processing. In this protocol, degradation is prevented by the addition of proteasome inhibitors during extraction, by performing the extraction on ice, and by fixing the cells rapidly after extraction.

Another key feature of the method is multiplexing, i.e., measurement of several different parameters simultaneously in single cells. One advantage of multiplexing is that it allows the incorporation of barcoding into the assay. In this protocol, barcoding is performed by labeling non-treated control cells with an amine-reactive Alexa 647, which binds to intracellular proteins (Krutzik and Nolan, 2006). The resulting Alexa 647–labeled cells are added to all the individual samples prior to staining with antibodies or the EdU Click-iT reaction and can be separated from the sample cells during analysis. The barcoded cells thus provide an internal standard for normalization of the RNAPII signals in the individual samples. This greatly facilitates quantification and accuracy of the assay and enables all the samples to be compared with each other. Multiplexing further permits analysis of RNAPII chromatin levels in specific phases of the cell cycle, by analyzing RNAPII chromatin levels only in cells with a certain DNA and EdU content (Bay et al., 2022).

Choice of antibodies is also critical in this method, as every new antibody used requires validation. The antibodies described here have been validated by us and others (Heidemann et al., 2013; Steurer et al., 2018; Bay et al., 2022), and recognize the N-terminal domain of RNAPII and the serine 5– (pRNAPII S5) and serine 2–phosphorylated (pRNAPII S2) forms of RNAPII. pRNAPII S5 is mostly associated with promoter-proximal regions, and global levels can therefore indicate degree of promoter-proximal paused RNAPII (Bay et al., 2022). pRNAPII S2 can be used as a marker for productive elongation (Bay et al., 2022). Finally, the use of EdU to label replicating cells is an important step to distinguish between G1 and early S phase, as well as between late S and G2. EdU is a nucleotide analogue, which is incorporated into the DNA molecule upon DNA replication. As their DNA content is highly similar, G1 vs. early S phase and G2 vs. late S-phase cells are not distinguishable by measurements of DNA content alone. By including EdU labeling, cells with similar DNA content can be separated in G1 vs. early S or G2 vs. late S based on the presence or absence of EdU incorporation, which marks the replicating (S phase) cells. However, this step may be omitted if such finely separated cell cycle determination is not required. If EdU is omitted, determination of cell cycle phase can be done by DNA content alone to obtain G1, S, and G2 fractions [Figure 2 and Dale Rein et al. (2015)].