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Co-phase separation of RNAs and RNA-binding proteins is thought to drive the biogenesis of ribonucleoprotein granules. RNAs can also undergo phase transitions in the absence of proteins. However, the physicochemical driving forces of protein-free, RNA-driven phase transitions remain unclear. Here, we report that RNAs of various types undergo phase transitions with system-specific lower critical solution temperatures (LCSTs). This entropically-driven phase behavior requires Mg2+ions and is an intrinsic feature of the phosphate backbone that is modulated by RNA bases. RNA-only condensates can additionally undergo enthalpically favorable percolation transitions within dense phases. This is enabled by a combination of Mg2+-dependent bridging interactions among phosphate groups and RNA base-stacking / base-pairing. Phase separation coupled to percolation can cause dynamical arrest of RNAs within condensates and can suppress the catalytic activity of an RNase P ribozyme. Our work highlights the need to incorporate RNA-driven phase transitions into models for RNP granule biogenesis.

Sensitive quantification of RNA transcripts via fluorescence in situ hybridization (FISH) is a ubiquitous part of understanding quantitative gene expression in single cells. Many techniques exist to identify and localize transcripts inside the cell, but often they are costly and labor intensive. Here we present a method to use a singly labeled short DNA oligo probe to perform FISH in yeast cells. This method is effective for highly constrained FISH applications where the target length is limited (200 nucleotides). This method can quantify different RNA isoforms or enable the use of fluorescence resonance energy transfer (FRET) to detect co-transcription of neighboring sequence blocks. Since this method relies on a single probe, it is also more cost-effective than a multiple probe labeling strategy.

Sensitive quantification of RNA transcripts via fluorescence in situ hybridization (FISH) is a ubiquitous part of understanding quantitative gene expression in single cells. Many techniques exist to identify and localize transcripts inside the cell, but often they are costly and labor intensive. Here we present a method to use a singly labeled short DNA oligo probe to perform FISH in yeast cells. This method is effective for highly constrained FISH applications where the target length is limited (

RNase P and MRP are highly conserved, multi-protein/RNA complexes with essential roles in processing ribosomal and tRNAs. Three proteins found in both complexes, Pop1, Pop6, and Pop7 are also telomerase-associated. Here, we determine how temperature sensitive POP1 and POP6 alleles affect yeast telomerase. At permissive temperatures, mutant Pop1/6 have little or no effect on cell growth, global protein levels, the abundance of Est1 and Est2 (telomerase proteins), and the processing of TLC1 (telomerase RNA). However, in pop mutants, TLC1 is more abundant, telomeres are short, and TLC1 accumulates in the cytoplasm. Although Est1/2 binding to TLC1 occurs at normal levels, Est1 (and hence Est3) binding is highly unstable. We propose that Pop-mediated stabilization of Est1 binding to TLC1 is a pre-requisite for formation and nuclear localization of the telomerase holoenzyme. Furthermore, Pop proteins affect TLC1 and the RNA subunits of RNase P/MRP in very different ways., Pop1 and 6 are subunits of RNase P and RNase MRP, which process ribosomal and tRNAs. The authors show that when Pop1 and 6 are impaired, the telomerase subunit Est1 binds telomerase RNA at normal levels, but the binding is unstable. As a result, nuclear import of the telomerase holoenzyme is inhibited.

mRNA Fluorescence In Situ Hybridization (FISH) is a technique commonly used to profile the distribution of transcripts in cells. When combined with the common single molecule technique Fluorescence Resonance Energy Transfer (FRET), FISH can also be used to profile the co-expression of nearby sequences in the transcript to measure processes such as alternate initiation or splicing variation of the transcript. Unlike in a conventional FISH method using multiple probes to target a single transcript, FRET is limited to the use of two probes labeled with matched dyes and requires the use of sensitized emission. Any widefield microscope capable of sensitive single molecule detection of Cy3 and Cy5 should be able to measure FRET in yeast cells. Alternatively, a FRET-FISH method can be used to unambiguously ascertain identity of the transcript without the use of a guide probe set used in other FISH techniques.

Quantitative profiling of mRNA expression is an important part of understanding the state of a cell. The technique of RNA Fluorescence In Situ Hybridization (FISH) involves targeting an RNA transcript with a set of 40 complementary fluorescently labeled DNA oligonucleotide probes. However, there are many circumstances such as transcripts shorter than 200 nt, splicing variations, or alternate initiation sites that create transcripts that would be indistinguishable to a set of multiple probes. To this end we adapted the standard FISH protocol to allow the use of a single probe with a single fluorophore to quantify the amount of transcripts inside budding yeast cells. In addition to allowing the quantification of short transcripts or short features of transcripts, this technique reduces the cost of performing FISH.

Quantitative measurement of mRNA levels in single cells is necessary to understand phenotypic variability within an otherwise isogenic population of cells. Single-molecule mRNA Fluorescence In Situ Hybridization (FISH) has been established as the standard method for this purpose, but current protocols require a long region of mRNA to be targeted by multiple DNA probes. Here, we introduce a new single-probe FISH protocol termed sFISH for budding yeast,Saccharomyces cerevisiaeusing a single DNA probe labeled with a single fluorophore. In sFISH, we markedly improved probe specificity and signal-to-background ratio by using methanol fixation and inclined laser illumination. We show that sFISH reports mRNA changes that correspond to protein levels and gene copy number. Using this new FISH protocol, we can detect more than 50% of the total target mRNA. We also demonstrate the versatility of sFISH using FRET detection and mRNA isoform profiling as examples. Our FISH protocol with single-fluorophore sensitivity significantly reduces cost and time compared to the conventional FISH protocols and opens up new opportunities to investigate small changes in RNA at the single cell level.

Quantitative determination of mRNA copy number distribution is essential for understanding phenotypic variability of single cells. Long mRNA transcripts (> 500 nt) can be detected using single-molecule Fluorescence In Situ Hybridization (FISH), where the target mRNA is hybridized with a large number of fluorescently labeled DNA probes. One downside to this method is the inability to interrogate short targets. We demonstrate a novel mRNA detection protocol based on Fluorescence Resonance Energy Transfer (FRET) which uses only a single pair of singly labeled DNA probes, each 24-nt long. Compared to conventional FISH, our method features higher signal-to-noise and lower cellular auto-fluorescence, thus effectively eliminating false positives. Using this technique in yeast, we estimate that ∼50% of transcripts can be detected. Our method will provide a reliable and efficient means to quantify relative levels of short mRNA transcripts in single cells.

Quantitative determination of the copy number of RNA transcripts in single cells is crucial to understand the genotype-phenotype connection. Current single-molecule Fluorescence In Situ Hybridization (FISH) protocols commonly used for this purpose require a large number of fluorophores per target RNA for single RNA detection, thus limiting the length range of RNA that can be probed. We have developed a FISH protocol for budding yeast, which can detect RNA molecules with a singly labeled 24-nucleotide DNA probe. Our single-probe protocol features highly inclined illumination and methanol fixation. We demonstrated high signal-to-noise and specificity of our protocol when tested against both constitutive and inducible genes in budding yeast. The technique presented offers a cost effective and efficient means of quantifying short RNA transcripts at the single cell level.