ARTICLE

DOI: 10.1038/s41467-018-06072-w OPEN Compartmentalised RNA catalysis in membrane-free

Björn Drobot1, Juan M. Iglesias-Artola 1, Kristian Le Vay 2, Viktoria Mayr2, Mrityunjoy Kar1, Moritz Kreysing 1, Hannes Mutschler 2 & T-Y Dora Tang 1

Phase separation of mixtures of oppositely charged provides a simple and direct route to compartmentalisation via complex coacervation, which may have been important for

1234567890():,; driving primitive reactions as part of the RNA world hypothesis. However, to date, RNA catalysis has not been reconciled with coacervation. Here we demonstrate that RNA catalysis is viable within coacervate microdroplets and further show that these membrane-free dro- plets can selectively retain longer length RNAs while permitting transfer of lower molecular weight oligonucleotides.

1 Max-Planck Institute for Molecular Biology and Genetics, Pfotenhauerstraße 108, 01307 Dresden, Germany. 2 Max-Planck Institute for Biochemistry, Am Klopferspitz 18, 82152 Martinsried, Germany. These authors contributed equally: Juan M. Iglesias-Artola, Kristian Le Vay. Correspondence and requests for materials should be addressed to M.K. (email: [email protected]) or to H.M. (email: [email protected]) or to T-Y.D.T. (email: [email protected])

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ompartmentalisation driven by spontaneous self-assembly ab processes is crucial for spatial localisation and con- 5′ C G centration of reactants in modern biology and may have G G BHQ1 been important during the origin of . One route known as G U 1,2 A complex coacervation describes a complexation process A G G C G C C G between two oppositely charged polymers such as polypeptides G C 3–7 U A (i)(ii) (iii) and nucleotides . The resulting coacervate microdroplets are A U G C membrane free, chemically enriched and in dynamic equilibrium C G A U G A A U with a poor phase. In addition to being stable over a A G broad range of physicochemical conditions, coacervate droplets A U C C

HH-min U are able to spatially localise and up-concentrate different AG A U

3,8 9,10 FRET-substrate molecules and support biochemical reactions . G C It has been hypothesised that compartments which form via G C C G coacervation could have played a crucial role during the origin of A U life by concentrating molecules and thus initiating the first bio- C chemical reactions on Earth11. Coacervation has also been A FAM 3′ implicated in modern biology where it has been shown that the formation of membrane-free compartments or condensates such c as P-bodies or stress granules within cells are driven by this (i) (ii) mechanism12,13. These membrane-free organelles are chemically 1 isolated from their surrounding cytoplasm through a diffusive 1 phase boundary, permitting the exchange of molecules with their 0.8 surroundings14. In addition, these compartments may localise 0.8 specific biological reactions and play important roles in cellular functions such as spatial and temporal RNA localisation within 0.6 0.6 the cell15–18. Whilst there is increasing evidence for the functional impor- 0.4 0.04 0.4 tance of RNA compartmentalisation via coacervation in modern Cleaved fraction 0 biology, this phenomenon would also have been vitally important 0.2 –0.05 0.2 during a more primitive biology. Up-concentration and locali- 02040 sation could have enabled RNA to function both as a catalyst 0 0 () and storage medium for genetic information, as 0 200 400 0 200 400 required by the RNA world hypothesis19. To date, have Time (min) Time (min) been encapsulated within eutectic ice phases20,21 and models such as water–oil-droplets for directed evolution experi- d ments22–24, membrane-bound lipid vesicles25–27, and membrane- (i) (ii) free compartments based on polyethelene glycol (PEG)/dextran 1 1 aqueous two-phase systems (ATPS)28. Interestingly, RNA cata- 0.01 lysis within ATPS exhibits an increased rate of reaction as a result 0.8 0 0.8 of the increased concentration within the dextran phase. Despite –0.02 0 400 these examples, RNA catalysis has not been demonstrated within 0.6 0.6 coacervate-based protocells. Therefore, coacervate protocells based on carboxymethyl dextran sodium salt (CM-Dex) and 0.4 0.4 poly-L-lysine (PLys) (Supplementary Fig. 1) were chosen as the model system due to their proven ability to encapsulate and Cleaved fraction support complex biochemical reactions catalysed by highly 0.2 0.2 evolved enzymes10. In contrast to these enzymes, structurally simple ribozymes, which are thought to have played a key role 0 0 during early biology, are prone to fold into inactive conforma- 0 200 400 0 200 400 tions in the absence of RNA chaperones or additional auxiliary Time (min) Time (min) elements29–33, and therefore may be rendered inactive by inter- actions within the highly charged and crowded interior of coa- cervate microdroplets. Herein, we directly probe the effect of the Methods) were incubated within a bulk polysaccharide/polypep- coacervate microenvironment on primitive RNA catalysis, and tide coacervate phase or within coacervate microdroplets under show the ability of the coacervate microenvironment to support single turnover conditions (see Methods). Cleavage of the FRET- RNA catalysis whilst selectively sequestering ribozymes and substrate strand by HH-min increases the distance between 6- permitting transfer of lower molecular weight oligonucleotides. carboxyfluorescein (FAM) and Black Hole quencher 1 (BHQ1), resulting in increased fluorescence intensity. We further devel- oped an inactive control ribozyme (HH-mut) by introducing two Results point mutations at the catalytic site (see Methods). Hammerhead activity in bulk coacervate phase. We developed a HH-min (1 µM) and FRET substrate (0.5 µM) were incubated real-time fluorescence resonance energy transfer (FRET) assay within the CM-Dex : PLys bulk coacervate phase (4:1 final molar (see Methods) to investigate the effect of the coacervate micro- concentration, pH 8.0). The RNA was then separated from the environment on catalysis of a minimal version of the hammer- coacervate phase and analysed by denaturing gel electrophoresis head ribozyme derived from RNA of tobacco ringspot (see Methods, Fig. 1). Excitingly, fluorescence gel imaging showed (HH-min)34. HH-min and its FRET substrate (Fig. 1a, the presence of cleavage product in the bulk coacervate phase

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Fig. 1 Cleavage of the FRET substrate under different conditions. a HH-min a (i) FAM-substrate (ii) TAM-HH-min (black) and the FRET substrate (red). b Gel electrophoresis of RNA cleavage in bulk coacervate phase (CM-Dex : PLys, 4:1 final molar ratio); μ μ 0.5 M of FRET substrate was incubated with 1 M of (i) HH-min, (ii) HH- –0.5 s mut or (iii) no ribozyme in bulk phase (25 °C, 60 min). Samples were analysed by denaturing PAGE followed by fluorescence imaging. The lack of in-gel quenching of the FRET substrate likely results from modifications of BHQ1 during PAGE51. c Real-time cleavage kinetics in 10 mM Tris-HCl pH

8.3 and 4 mM MgCl2. (i) A monoexponential fit (Methods, Eq. 3) (grey line) to kinetic data (grey dots) and residuals of the fit (inset); (ii) mean of 0 s the individual fits to each experiment (Blue line) with the standard deviation of the mean of the fits (grey data points) (N = 5). d Cleavage in bulk coacervate phase (normalised to the amount of cleaved product at t = 530 min from gel electrophoresis). (i) Biexponential fit (Methods, Eq. 4) (dark grey line) to experimental data (grey dots) with the residuals (inset); (ii) 13 s mean biexponential fit (orange) of individual fits (N ≥ 5). Grey data points represent the standard deviation (N = 5) from the experimental data

b (i) (ii) 1 1 containing HH-min (Fig. 1b). In contrast, control experiments in 0.8 0.8 0.09 0.07 the absence of HH-min or in the presence of HH-mut showed no norm norm fi I 0.6 0 I 0.6 0 evidence of the cleavage product, con rming that the wild-type –0.08 –0.05 ribozyme drives substrate cleavage in the bulk coacervate phase 0.4 0510 0.4 0510 (Fig. 1b). The FRET assay was further exploited to characterise 0510 0510 the enzyme kinetics in both the bulk coacervate phase and buffer Time (s) Time (s) fl by time-resolved uorescence spectroscopy under single turnover c 150 conditions by direct loading of HH-min and FRET substrate into 1.5 either cleavage buffer or bulk coacervate phase (see Methods).

/s) 100 The increase in fluorescence intensity of FAM was measured over 2 1 m μ (mPa s) time and normalised to the amount of cleaved product generated. ( 50 D Fitting the kinetic profiles of substrate cleavage in buffer 0.5  conditions with a single exponential revealed an apparent rate 0 0 = (i) (ii) constant, k0 of 0.6 ± 0.1/min (Fig. 1c, N 5), which was comparable to the k0 obtained in buffer analysed by gel Fig. 2 FRAP of bulk coacervate phase. Bulk coacervate phase (CM-Dex:PLys electrophoresis (0.38 ± 0.05/min) (see Methods, Supplementary fi μ = (4:1 nal molar ratio) containing either (i) 0.36 M FAM-substrate or (ii) Fig. 2, N 6) and to kcat values previously determined for a range 0.36 μM TAM-HH-min. a Output frames from confocal imaging (63×) are – 34,35 of hammerhead ribozymes (0.01 1.5/min) . In comparison, shown at t = −0.5 s before bleaching, directly after bleaching (magenta RNA cleavage within the bulk coacervate phase was clearly circle, t = 0 s) and t = 13 s after bleaching. The fluorescence intensity was biphasic (Supplementary Fig. 3A) with an observed faster rate normalised against a reference (green circle) and fit to standard equations. –2 –2 constant, k1, of 1.0 × 10 ± 0.1 × 10 /min and a second slower Scale bars are 5 μM. b Plots of normalised FRAP data for HH-min (ii) and –4 –4 rate constant, k2, 1.5 × 10 ± 0.8 × 10 /min (errors obtained FAM-substrate (ii) show the standard deviation (grey, N = 10) and fit from 12 individual droplets). Thus, the fastest rate constant k1 is fi = (blue) from the same bleach spot radius. c Diffusion coef cients and 60-fold slower than in buffer conditions (k0 0.6 ± 0.1/min) viscosities obtained from b. Mean and standard deviations are from at least indicative of reduced activity within the coacervate phase. In two different samples with analysis from ≥14 bleach spots for each addition, the transition to biphasic kinetics within the coacervate experiment phase compared to the aqueous buffer phase describes a different kinetic mechanism of HH-min within the coacervate phase deviations are from at least two different samples with analysis (Fig. 1d). This may be attributable to heterogeneous ribozyme from at least 14 bleach spots from each experiment) from populations with variations to secondary structure and/or Fluoresence Recovery after Photobleaching (FRAP) analysis alternative conformational and equilibrium states, as observed showed a significantly slower molecular diffusion of the ribozyme for some HH systems in aqueous buffer conditions36,37. Circular fi θ and substrate compared to predicted diffusion coef cients of dichroism (CD) spectra show a reduction in molar ellipticity ([ ]) RNA in buffer (~150 µm2/s, Fig. 2)38,39. The decreased mobility is for HH-mut within bulk coacervate phase compared to aqueous indicative of a more viscous and spatially restricted environment buffer with a small commensurate shift in the peak maxima from in the interior of the coacervate phase (η = 114 ± 21 mPa. s, 265 nm to 268 nm, respectively (Supplementary Fig. 4). These Fig. 2c). results show that the fold of HH-mut is altered in the polyelectrolyte-rich bulk coacervate phase, with an overall loss of secondary structure that could affect catalytic activity. In Hammerhead activity in coacervate microdroplets. To test the addition, it is possible that the charged and crowded coacervate activity of the ribozyme within individual droplets, the bulk microenvironment restricts substrate binding, sterically hinders coacervate phase containing ribozyme and substrate was re- substrate–enzyme complex formation and/or spatially restricts dispersed in supernatant to produce microdroplets in solution diffusion of the cleavage assay components. Indeed, measured (see Methods). The final concentration of enzyme and substrate diffusion coefficients of HH-min tagged with TAMRA (TAM- in the microdroplet dispersion, formed from 1 µl of bulk coa- HH-min) (1.0 ± 0.2 µm2/s) and FAM-substrate (1.6 ± 0.1 µm2/s) cervate phase redispersed in 49 µl of supernatant was equivalent in the bulk coacervate phase (Fig. 2, mean and standard to the final concentration of the bulk coacervate phase i.e. within

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ab (i) (ii) (iii)

×103 ×103 30 30 1 Coacervate microdroplets 20 20 10 10 0 0

Intensity (a.u.) 0 5 10 15 Intensity (a.u.) 0 5 10 15 Length (μm) Length (μm)

0.5 0.05 0 –0.05 Cleaved fraction 0 200 400 600 800 0 0 200 400 600 800 Time (min) 0 min0 min 900 min

Fig. 3 RNA catalysis in coacervate microdroplets. a (i) Wide-field optical microscopy images of CM-Dex:PLys (4:1 final molar ratio) coacervate microdroplets prepared in cleavage buffer (1 μM of HH-min and 0.5 μM FRET substrate). Fluorescence microscopy images at t = 0 min (ii) and t = 900 min (iii) show an increase in FAM fluorescence (see inset). Scale bars are 20 μM. b Background corrected and volume/endpoint normalised fluorescence intensity of droplets. Standard deviation of kinetics from 12 micro-droplets (grey) with the mean biexponential fit (blue) and residuals (inset)

50 µl of phase under single turnover conditions (1 µM of HH-min Selective RNA partitioning into . To further inves- and 0.5 µM FRET substrate). Fluorescence optical microscopy tigate the six-fold difference in the partition coefficient between images showed an increase in FAM fluorescence intensity in the the ribozyme and substrate, we characterised the differences in droplets after 900 min (Fig. 3a). the rate and extent of sequestration of TAM-HH-min and FAM- Fitting the biphasic fluorescence signal allowed a direct substrate from the surrounding aqueous phase into the droplet comparison of kinetic profiles between the coacervate micro- after whole-droplet photobleaching. Coacervates containing droplet (Fig. 3b, Supplementary Fig. 3B) and bulk coacervate FAM-substrate (12-mer) showed complete fluorescence recovery phase environments. A modestly faster rate constant (k1 and k2) within 100 s and a recovery half time (τ) of 22 ± 3.5 s (N = 20). In –2 –2 was observed in the microdroplets (k1 of 4.4 × 10 ± 1.3 × 10 / comparison, TAM-HH-min showed only 70% recovery after 300 –3 –3 min, k2 of 2.3 × 10 ± 0.2 × 10 /min, N = 12) compared to the s with τ = 189 ± 14 s (N = 11), attributed to either a low con- –2 –2 bulk coacervate phase (k1 of 1.0 × 10 ± 0.1 × 10 /min, k2 of centration of HH-min within the surrounding aqueous phase, a 1.5 × 10–4 ± 0.8 × 10–4/min, N = 11) (Supplementary Table 3, slow rate of transfer into the coacervate droplet from its exterior Supplementary Fig. 5). Determination of the partition coefficients and/or immobilised ribozyme in the coacervate droplet, which is of both the ribozyme and substrate (KHH-min = 9600 ± 5600 unable to exchange with RNA in the surrounding aqueous phase = = = (N 12) and KHH-substrate 3000 ± 2000 (N 20), Supplemen- (Supplementary Fig. 8). The results from the FRAP experiments tary Fig. 6) by fluorescence spectroscopy (see Methods) showed complement the equilibrium partition coefficient. The 12-mer that both TAM-HH-min and FAM-substrate partition strongly substrate, with a lower partition coefficient (K = 3000 ± 2000, into the coacervate environment. 1 μl of bulk coacervate phase N = 20) shows a faster exchange between the droplet and was used to prepare coacervate microdroplet suspensions in a the surrounding aqueous phase compared with the 39-mer total volume of 50 μl compared to 50 μl of bulk coacervate phase ribozyme, which has a higher partition coefficient (K = 9600 for bulk experiments. Thus, based on the measured partition ± 5600 (N = 12)), shows slower exchange of RNA with the sur- coefficients, we calculated concentrations of 49.6 μM HH-min rounding environment. A strong correlation between FRAP half- and 24.3 μM substrate in a single microdroplet, compared to and partitioning coefficients was also described for RNAs in 1 μM HH-min and 0.5 μM substrate within the bulk coacervate other coacervate systems4. phase. Whilst the observed rate differences between the bulk To investigate additional sequence-dependent effects on coacervate and coacervate microdroplet phases could be due to partitioning, we compared the partition coefficients of different variations in viscosity, quantitative FRAP analysis with two 12-mer RNAs (Supplementary Fig. 6): FAM-substrate is pyr- different FAM-substrate concentrations (0.36 μM and 24.3 μM) imidine rich but unstructured RNA; FAM-tet is a pyrimidine-rich showed that the measured viscosities of the bulk coacervate phase hairpin structure with a stable UUCG tetraloop; FAM-flex is an and coacervate microdroplet are comparable within error unstructured purine-rich sequence (see Supplementary Table 2). (Supplementary Fig. 7). Therefore, secondary effects arising from Our results show that for unstructured RNAs an increase in the increased RNA concentration within the coacervate micro- purine content reduces partitioning of 12-mer RNAs (FAM- droplet phase may be responsible for the increased rate constants substrate vs. FAM-flex) approximately 10-fold. Likewise, we observed. The ribozyme and substrate concentrations are observe a decrease in the partition coefficient with an increase in approximately 50 times more in the coacervate microdroplet secondary structure for pyrimidine-rich RNA (FAM-substrate vs. (49.6 μM and 24.3 μM, respectively) compared to the bulk FAM-tet) (Supplementary Fig. 9). Thus, our results, and coacervate phase (1 μM and 0.5 μM, respectively). The difference others4,42,43, show that RNA sequestration and localisation in diffusion length scales of RNA in the microdroplet environ- within coacervate droplets is dependent on the length, sequence ment could lead to increased saturation of the ribozyme and and also structure of the sequestered RNA. therefore greater apparent rate constants in the coacervate For the RNAs specific to our HH ribozyme assay, the general microdroplets compared to bulk coacervate phase. In addition, selective retention of longer length polynucleotides with transfer enrichment of RNA could lead to changes in material properties of shorter length RNA can have interesting implications for such as water activity or dielectric constant, which have a direct ribozyme catalysis within coacervate droplets. To investigate this, impact on the rate of hammerhead-catalysed RNA cleavage40,41. we directly observed the localisation of TAM-HH-min and Thus, secondary effects that result from the increased RNA FRET-substrate. CM-Dex:PLys coacervate micro-droplets (4:1 concentration within coacervate microdroplets may concomi- final molar ratio) containing TAM-HH-min were loaded into one tantly contribute to an increase in the apparent rate constant. end of a capillary channel (Fig. 4a, region 1) while droplets

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a b

Microscope channel

TAM-HH-min 1 1

2

3 FRET substrate

2 c FAM channel

150 1

100

50 2 3 Intensity (a.u.) 3 0 0 200 400 Time (min) (i) Brightfield (ii) TAM channel (iii) FAM channel

Fig. 4 Localisation and retention of RNA within coacervate droplets. a Schematic of localisation experiments where CM-Dex:PLys (4:1 final molar ratio) coacervate droplets containing 0.36 μM(final concentration) TAM-HH-min were loaded into one end of a capillary channel (1). Droplets containing 0.36 μM FRET-substrate were loaded into the other end of the channel (3). b Wide-field optical microscopy images obtained using a 100 × oil immersion lens in (i) bright field and fluorescence mode using filters for (ii) TAM or (iii) FAM. Images were captured in regions 1, 2 and 3 at t = 500 min (scale bar: 20 μM). c FAM fluorescence intensity. Shaded regions represent the standard deviation of at least seven droplets containing the FRET-substrate were loaded into the other end of microdroplets (4:1 final molar ratio) at pH 8: droplets containing the channel (Fig. 4a, region 3) in such a way as to prevent droplet either TAM-HH min or FAM-substrate were loaded into one end mixing whilst permitting passive diffusion of molecules through of a capillary channel, and coacervate droplets containing HH- the length of the channel (see Methods). Time-resolved mut were loaded into the other end of the capillary channel in fluorescence optical microscopy images in both the TAM and such a way as to prevent droplet mixing (Supplementary Fig. 11). FAM channels were obtained at various locations along the Fluorescence optical microscopy images obtained in the middle of capillary channel (Fig. 4a, regions 1, 2, 3). Imaging over 500 min the channel (Supplementary Fig. 11B, region 2) showed no in the TAM channel showed that, within the measurable change in the fluorescence intensity of TAM-HH-min (Supple- resolution, no diffusion of the ribozyme to droplets in other mentary Fig. 11C, i) over the course of the experiment (500 min). regions of the channel occurs (Fig. 4b, region 2 and Supplemen- In contrast, a small increase in the fluorescence intensity from tary Fig. 10). Conversely, over the time course, droplets in all FAM-substrate (Supplementary Fig. 11C, ii) was observed after three regions exhibited increased FAM fluorescence with droplets 300 min, suggesting a higher exchange rate of the 12-mer with the in region 1 with the highest intensity and droplets in regions 2 environment. Both the partition coefficient and whole droplet and 3 with comparatively lower intensity (Fig. 4c). Analysis of FRAP results for PLys:ATP coacervate microdroplets, containing time-resolved images showed a delayed increase in the onset of either TAM-HH-min (39-mer), FAM-substrate (12-mer) or cleaved product fluorescence in region 2 and a further delayed cleaved FAM-substrate (6-mer) (Supplementary Figs. 12, 13), onset in cleaved product in region 3. These results are confirmed a consistent trend in RNA retention based on RNA commensurate with diffusion of the FRET-substrate out of the length with an order of magnitude difference in τ between the droplets in region 3 and into droplets containing TAM-HH-min different oligonucleotides (Supplementary Fig. 12). Whilst the in region 1 where cleavage takes place. The cleaved product then general trends are consistent with those observed with CM-Dex: diffuses out of the active droplets and into droplets in regions 2 PLys coacervate microdroplets, a direct comparison of whole and 3. Control experiments probing the transfer of FAM- droplet FRAP recovery times (Supplementary Table 4) between substrate only show increased fluorescence intensity in regions 2 the two microenvironments shows that RNA has a stronger and 3 giving a direct confirmation that the 12-mer substrate tendency to localise within the PLys:ATP microdroplets com- diffuses between droplets (Supplementary Fig. 10). Taken pared to CM-Dex:PLys microdroplets. The longer recovery times together, our results show that longer length RNA (HH-min) is after whole droplet photobleaching in the PLys:ATP droplets retained and spatially localised within the highly charged and could be attributed to differences in molecular interactions of crowded interior of the coacervate droplet, while shorter RNAs RNA within the two microenvironments. For example, there may transfer between the droplets. be increased RNA–PLys interaction in the PLys:ATP environ- As other studies have shown that RNA rapidly exchanges from ment compared to the PLys:CM-Dex, where presence of CM-Dex PLys and adenosine triphosphate (ATP, Supplementary Fig. 1) could shield PLys–RNA interactions. In addition, favourable coacervate microdroplets into the surrounding environment44,we Pi–Pi stacking interactions between the aromatic rings of ATP also tested this coacervate system for selective localisation of and RNA would favour interactions of RNA within the PLys:ATP RNA. To this end, localisation experiments were undertaken as system. Taken together, the results show that membrane-free previously described (see Methods) with PLys:ATP coacervate droplets prepared via coacervation offer general features such as

NATURE COMMUNICATIONS | (2018) 9:3643 | DOI: 10.1038/s41467-018-06072-w | www.nature.com/naturecommunications 5 ARTICLE NATURE COMMUNICATIONS | DOI: 10.1038/s41467-018-06072-w length-dependent RNA localisation and transfer. Moreover, our site protonation−deprotonation events, respectively46. The wild-type and inactive results also show that the strength of oligonucleotide selectivity is hammerhead ribozymes were transcribed in vitro by T7 RNA polymerase. The fi dependent on the composition of the coacervate microdroplets DNA templates for were produced by ll-in of DNA oligonucleotides using GoTaq (Promega). Complementary pairs of DNA oligonucleotides contained and the molecular sequence of RNA. the ribozyme (sTRSV_min_wt_TX/ sTRSV_min_mut_TX) and T7 promoter (5T7) (Supplementary Table 1). The double-stranded templates were purified using Discussion a Monarch PCR DNA Cleanup Kit (NEB, Biolabs, USA). Ribozyme RNA was transcribed from the double-stranded templates using the MEGAshortscript™ T7 Here, we demonstrate that coacervate microdroplets offer intri- Transcription Kit (ThermoFisher), and purified using RNeasy (Qiagen). guing properties for compartmentalised RNA catalysis. Our results show that these membrane-free microenvironments sup- Preparation of coacervates containing RNA HH and substrate. Preparation of port RNA catalysis and up-concentrate oligonucleotides within bulk coacervate phase and coacervate microdroplets containing RNA HH and their interiors. This effect is coupled to selective retention and substrate. Aqueous dispersions of CM-Dex:PLys coacervate microdroplets (4:1 release of RNA without additional energy input. These features molar ratio) or ATP:PLys (4:1 molar ratio) in 10 mM Tris-HCl pH 8.0 and 4 mM MgCl were first prepared by mixing 200 µl of 1 M CM-Dex, 250 µl of 0.2 M PLys, could have been significant on where concentrations 2 50 µl of 1 M Tris-HCl pH 8.0 and 20 µl of 1 M MgCl2 and made up to 5 ml with of RNA and their building blocks may have been low. Moreover, nuclease-free water. Aqueous dispersions of PLys:ATP coacervate dispersions (4:1 maintenance of the genetic identity of coacervate protocells could molar ratio) in 10 mM Tris-HCl pH 8.0 and 4 mM MgCl2 were produced by be achieved via spatial localisation of RNA catalysis and RNA mixing 1000 µl of 0.2 M PLys, 500 µl of 0.1 M ATP, 50 µl of 1 M Tris-HCl pH 8.0 and 20 µl of 1 M MgCl2 and made up to 5 ml with nuclease-free water. To produce with spread of RNA building blocks or short genetic a polymer-only bulk coacervate phase (approximately 100 µl), the aqueous dis- polymers between droplets. Whilst this work represents a key step persion of microdroplets was centrifuged (10 min, 4000×g) and the supernatant in reconciling primitive RNA catalysis with selective protocellular removed. To produce bulk coacervate phase containing RNA, RNA was added compartmentalisation, it should also be noted that these features directly to 80 µl of bulk coacervate phase. The samples were mechanically mixed fi and centrifuged (10 min, 4000×g) and any excess water from the RNA stock of compartmentalisation are signi cant in modern biology. To solutions was removed. To produce coacervate microdroplet dispersions contain- this end, there are immediate questions to be addressed. For ing the same total concentration of RNA compared to bulk coacervate phase, 1 µl example, how does the microdroplet environment alter the of bulk coacervate phase loaded with RNA (e.g., 50 pmol HH-min) was prepared as ribozyme mechanistic pathway and effect nucleotide selectivity previously described. This 1 µl of the bulk coacervate loaded with RNA was made up to 50 µl with supernatant and vortexed to produce a dispersion of RNA loaded into the droplet? How can we alter the physical chemistry of the coacervate microdroplets. Final concentrations of RNA are calculated from the droplets to further affect oligonucleotide selectivity? In addition, total volume, i.e., bulk coacervate samples contain 50 μl of bulk coacervate phase our experiments have focused on a nucleolytic ribozyme; how- whilst the volume of the microdroplet dispersions accounts for both the volume of ever, rather than RNA cleavage, an increase in genetic and coacervate bulk phase and the supernatant it is dispersed in. Therefore, whilst the molecular complexity of RNA, e.g., by ligation would have been total concentrations are equivalent, there is approximately 50× less coacervate phase within the dispersion (1 μl) compared to the bulk coacervate phase important during early Earth. Therefore, probing RNA synthesis (50 μl). through ligase activity within coacervate microdroplets will fur- ther contribute to the understanding of the role of coacervation Hammerhead ribozyme FRET assay. A minimal trans-cleaving version of the on early Earth and modern biology. tobacco ring spot virus hammerhead ribozyme (HH-min) was chosen as the model system for this study. To characterise the activity of HH-min, we employed a FRET assay. Cleavage of the FRET-substrate strand by HH-min increases the distance Methods between FAM and BHQ1, resulting in increased fluorescence intensity. In addition, Materials. Trizma base (Tris), CM-Dex sodium salt (10–20 kDa, monomer: a modified version of the HH-min was developed as an inactive control ribozyme 162.14 g/mol), PLys hydrobromide (4–15 kDa, monomer: 161.67 g/mol), (HH-mut) by introducing two point mutations at the catalytic site. This mutant ATP disodium salt (551.14 g/mol), sodium hexametaphosphate ((NaPO3)6, permits binding of the substrate, but is catalytically inactive. 611.77 g/mol), formamide (CH3NO, 45.04 g/mol), ethylenediaminetetracetic acid disodium salt dihydrate (EDTA, C H N Na O ·2H O, 372.24 g/mol), fluorescein 10 14 2 2 8 2 Hammerhead activity with gel electrophoresis. Substrate cleavage assays were isothiocyanate isomer I (FITC, C21H11NO5S, 389.38 g/mol), bromophenol blue −1 carried out in cleavage buffer (10 mM Tris-HCl, pH 8.3, and 4 mM MgCl2)at25°C (C19H10Br4O5S, 669.96 g mol ) and Orange G (C16H10N2Na2O7S2, 452.37 g/mol were purchased from Sigma Aldrich and used without further purification. Mag- under single turnover conditions, i.e., with stoichiometric amounts of ribozyme and substrate; in this case, the ribozyme concentration is 2× the concentration of nesium chloride hexahydrate (MgCl2·6 H2O, 203.30 g/mol), sodium hydroxide μ μ (NaOH, 39.997 g/mol), sodium chloride (NaCl, 58.44 g/mol), urea (CH N O, substrate. An excess of HH-min (1 M) compared to FRET-substrate (0.5 M) 4 2 were mixed in RNAse-free water, the reaction was initiated by the addition of 60.06 g/mol), hydrochloric acid (HCl, 37%, 36.46 g/mol) and boric acid (H3BO3, 61.83 g/mol) were purchased from Merck and used without further purification. cleavage buffer. Aliquots were taken at varying time points and quenched on ice by ® the addition of four volumes of RNA gel loading buffer (formamide, EDTA (10 Ammoniumperoxodisulfate (APS, (NH4)2S2O8, 228.20 g/mol), Rotiphorese Gel 40 (Acrylamide 37, 5:1 Bisacrylamide) and Tetramethylethylendiamine (TEMED, mM, pH 8.0), bromophenol blue (0.025% w/v). The substrate and cleavage pro- C H N , 116.21 g/mol) were purchased from Roth. 5(6)-Carboxyte- ducts were separated by denaturing PAGE on 20% acrylamide gels and run in 1× 6 16 2 TBE buffer. Bands were visualised by FAM-tag fluorescence (Typhoon FLA-5000, tramethylrhodamine succinimidyl ester (TAMRA, C29H25N3O7, 527.53 g/mol) and λ = λ = SYBR gold stain (10,000× concentrate in DMSO) were purchased from Thermo GE Healthcare Life Sciences, ex 473 nm, em 520 nm), and the intensities of Fisher Scientific. Oligo length standards (10/60 single strand RNA) was purchased cleavage product at each time point were determined by integration of band from Integrated DNA technologies. intensities using ImageQuant. fi The first-order time constant τ was obtained by fitting the cleaved and All DNA and tagged RNA oligonucleotides were synthesised by Euro ns fi fi Ebersberg, Germany, and used without further purification (Supplementary uncleaved kinetic pro les to single exponential ts (Eqs. 1, 2, respectively), which Table 2). Stocks of the RNA constructs were prepared in nuclease-free water and were globally optimised. stored at −80 °C until use. ÀðÞt ð Þcleaved ¼ cleaved À cleaved ´ τ þ ; ð1Þ Nuclease-free water was purchased from Ambion and used to prepare aqueous I t Imax Imax e C stock solutions of CM-Dex (1 M, pH 8), PLys (0.2 M, pH 8), ATP (0.1 M, pH 8) and stored in the freezer (−20 °C) until use. The pH of the stock solutions was ÀÁ ÀðÞt ð Þuncleaved ¼ þ uncleaved À ´ τ : ð Þ adjusted using NaOH, and concentrations were determined from the molecular I t C Imax C e 2 weight of the monomer. Aqueous stock solutions of Tris-HCl (1 M, pH 8/pH 8.3) and MgCl2 (1 M) were prepared and used for buffer solutions. Capillary channel slides for microscopy were custom made using PEGylated cover slips (22 × 22 mm) I(t) is the band intensity, C is an offset and τ the first-order time constant. The fi τ and microscope slides (25 mm × 75 mm). rst-order rate constant (k0) is then determined by 1/ . To obtain the differences in fluorescence quantum yield for the cleaved and uncleaved substrate (FRET effect), the maximum intensities at the start and the end RNA synthesis. A minimal, trans-acting hammerhead ribozyme (HH-min) points (I ) were obtained and their ratio determined. derived from satellite RNA of tobacco ringspot virus and complementary substrate max were produced by modification of the helix 1 hybridising arm in a cis-acting system45. An inactive variant (HH-mut) was produced by the introduction of two Gel electrophoresis of hammerhead activity in coacervates. HH-min and point mutations, G5A and G12A, which inhibit correct ribozyme folding and active FRET-substrate at final concentrations of 1 µM and 0.5 µM, respectively, were

6 NATURE COMMUNICATIONS | (2018) 9:3643 | DOI: 10.1038/s41467-018-06072-w | www.nature.com/naturecommunications NATURE COMMUNICATIONS | DOI: 10.1038/s41467-018-06072-w ARTICLE loaded into CM-Dex:PLys (4:1 final molar ratio) bulk coacervate phase for single microdroplets (4:1 final molar ratio) and the bulk coacervate phase (CM-Dex:PLys turnover conditions, i.e., with stoichiometric amounts of ribozyme and substrate; in 4:1 final molar ratio) containing either TAM-HH-min (0.36 µM), FAM-substrate this case, the ribozyme concentration is 2× the concentration of substrate. RNA (0.36 µM), FAM-substrate (24.3 µM) or FAM-cleaved substrate (0.36 µM). Samples was extracted from 5 µl of CM-Dex:PLys (4:1 final molar ratio) bulk coacervate were prepared as previously described and loaded into capillary slides mounted in a phase (25 °C) or from 5 µl of CM-Dex:PLys (4:1 final molar ratio) coacervate Zeiss LSM 880 inverted single point scanning confocal microscope equipped with a microdroplet dispersions after 900 min (25 °C) by sequential addition of 5 µl 1 M 32 GaAsP PMT channel spectral detector and a 32-channel Airy Scan detector and NaCl (final concentration 4.8 mM), 5 µl of 1.25 M hexametaphosphate (final imaged using a 63× oil immersion objective (Plan-Apochromat 63× 1.4 Oil DIC, concentration −6.0 mM) and 90 µl RNA loading buffer (final volume—83%) Zeiss). Bleaching was achieved by additional excitation with a 405 nm laser diode containing EDTA (final concentration—8 mM) and Orange G and 10 s of vor- and 355 nm DPSS laser, an Argon Multiline Laser produced the excitation wave- λ = λ = λ = texing and 1 s of centrifugation between each addition. The reaction mixture was length of FAM 488 nm or TAM 561 nm and emission wavelengths FAM – λ = – heated at 80 °C for 10 min, centrifuged and placed on ice for at least 5 min. 10 µl of 479 665 nm (Laser line blocking pin at 488 nm) or TAM 562 722 nm. Imaging the reaction mixture was loaded into a pre-run 20% polyacrylamide gel and then time varied depending on the region of interest but was typically between 12 ms/ run at 300 V in 1× TBE buffer until the dye had run to the bottom of the gel. The frame and 100 ms/frame. λ = fl gel was imaged using Typhoon FLA-9500, GE Healthcare Life Sciences, with ex The uorescence intensity as a function of time for the bleached area, reference λ = 473 nm and em 520 nm. Band intensities were measured using ImageQuant at a and the background was obtained using FIJI and the recovery of the bleached specific time point and uncleaved substrate was corrected by the FRET effect factor region was normalised against the background and the reference region for either (1.69) (Supplementary Fig. 12). The fraction of cleaved substrate of the total sum of bulk CM-Dex:PLys (4:1 final molar ratio) coacervate or PLys:ATP (4:1 final molar cleaved and uncleaved substrate was determined and used to normalise kinetic data ratio) coacervate experiments. An additional normalisation for droplet-based obtained from spectroscopy or microscopy at specific time points. Gel electro- FRAP was undertaken by dividing the fluorescence by the fluorescence of the whole phoresis was undertaken with FRET-substrate and FAM-substrate with a single- droplet47,48. The kinetic profiles were fit to Eq. 5 using MATLAB to obtain the time strand RNA molecular marker on a pre-run 20% polyacrylamide gel and run at 15 constant, τ,offluorescence recovery or of transport into the droplet (whole droplet W in 1× TBE buffer. The gel was stained with SYBR gold and imaged using a FRAP). λ = λ = Typhoon FLA-5000, GE Healthcare Life Sciences, with ex 473 nm and em ( ¼ 520 nm. Uncropped gels are shown in Supplementary Figure 15. 1 for t

NATURE COMMUNICATIONS | (2018) 9:3643 | DOI: 10.1038/s41467-018-06072-w | www.nature.com/naturecommunications 7 ARTICLE NATURE COMMUNICATIONS | DOI: 10.1038/s41467-018-06072-w

Fluorescence-based partitioning of RNA. TAM-HH-min, FAM-substrate, FAM- 2. Priftis, D., Laugel, N. & Tirrell, M. Thermodynamic characterization of cleaved product, FAM-Tet, FAM-Flex, FITC and TAMRA were loaded into 150 µl polypeptide complex coacervation. Langmuir 28, 15947–15957 dispersions containing 3 µl of bulk coacervate phase (CM-Dex:PLys 4:1 molar (2012). μ ratio) to achieve an initial concentration of 0.5 M(cini). After an equilibration 3. Koga, S., Williams, D. S., Perriman, A. W. & Mann, S. Peptide–nucleotide time of 10 min, the coacervate phase (3 µl) was separated from the supernatant microdroplets as a step towards a membrane-free protocell model. Nat. Chem. (147 µl) by centrifugation (10 min at 10,000×g). To ensure both fractions were 3, 720–724 (2011). treated equally, the 147 µl supernatant fraction was filled to a final volume of 150 µl 4. Aumiller, W. M., Pir Cakmak, F., Davis, B. W. & Keating, C. D. RNA-based fi (Vini) by the addition of 3 µl of RNA-free bulk CM-Dex:PLys (4:1 nal molar ratio) coacervates as a model for membraneless organelles: formation, properties, coacervate phase (supernatant phase), whilst the 3 µl of RNA-containing bulk 4:1 and interfacial liposome assembly. Langmuir 32, 10042–10053 (2016). fi CM-Dex:PLys (4:1 nal molar ratio) coacervate phase fraction was made up to 5. Priftis, D. & Tirrell, M. Phase behaviour and complex coacervation of aqueous 150 µl (Vini) with RNA-free supernatant (coacervate phase). Coacervates of both polypeptide solutions. Soft Matter 8, 9396–9405 (2012). samples were dissolved by the addition of 10 µl NaCl (5 M). The Fraction, F,of 6. de Kruif, C. G., Weinbreck, F. & de Vries, R. Complex coacervation of fi RNA within either the bulk CM-Dex:PLys (4:1 nal molar ratio) coacervate phase and anionic polysaccharides. Curr. Opin. Interface Sci. 9, 340–349 or the supernatant phase was obtained using Eq. 9: (2004). – Iphase 7. Vieregg, J. R. et al. Oligonucleotide peptide complexes: phase control by F ¼ P ; ð9Þ – phase I hybridization. J. Am. Chem. Soc. 140, 1632 1638 (2018). 8. Tang, T.-Y. D. et al. Fatty acid membrane assembly on coacervate where the I is the fluorescence intensity of either the coacervate or supernatant microdroplets as a step towards a hybrid protocell model. Nat. Chem. 6, phase – phase measured from 20 μl of sample using the TECAN spark 20M of either FAM- 527 533 (2014). λFAM = λFAM = λTAM = 9. Crosby, J. et al. Stabilization and enhanced reactivity of actinorhodin substrate ( exc 485 nm, em 535 nm) or TAM-HH-min ( exc 544 nm, λTAM= ∑ polyketide synthase minimal complex in polymer–nucleotide coacervate em 576 nm) and I is the sum of both. The corresponding concentrations in both phases were calculated using Eq. 10: droplets. Chem. Commun. 48, 11832–11834 (2012). 10. Tang, T.-Y. D., van Swaay, D., deMello, A., Anderson, J. L. R. & Mann, S. 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Biol. 30,39 58 (2014). 15. Banani, S. F., Lee, H. O., Hyman, A. A. & Rosen, M. K. Biomolecular where the c is the concentration of RNA in the bulk coacervate phase and c condensates: organizers of cellular biochemistry. Nat. Rev. Mol. Cell Biol. 18, coac super – is the concentration of RNA in the supernatant phase. 285 298 (2017). 16. Zhang, X. et al. RNA stores tau reversibly in complex coacervates. PLoS Biol. 15, e2002183 (2017). Ribozyme activity in supernatant phase . In order to estimate the contribution of 17. Shin, Y. & Brangwynne, C. P. Liquid phase condensation in cell physiology FRET-substrate cleavage in the aqueous surrounding of the coacervate micro- and disease. Science 357, eaaf4382 (2017). droplets, cleavage experiments were performed in the supernatant using a HH-min 18. Saha, S. et al. Polar positioning of phase-separated liquid compartments in concentration of 0.007 µM (based on the partitioning coefficient for the ribozyme) cells regulated by an mRNA competition mechanism. Cell 166, 1572–1584.e16 and 0.5 µM FRET-substrate concentration (maximum possible concentration). The (2016). data were compared to a control in supernatant under single turnover conditions 19. Joyce, G. F. The antiquity of RNA-based evolution. Nature 418, 214–221 (1 µM HH-min, 0.5 µM FRET-substrate). The kinetics of the substrate cleavage was obtained using TECAN Spark 20M well plate reader spectrophotometer (Tecan (2002). AG, Männedorf, Switzerland) by measuring the increase in FAM fluorescence over 20. Attwater, J., Wochner, A., Pinheiro, V. B., Coulson, A. & Holliger, P. Ice as a λ = λ = protocellular medium for RNA replication. Nat. Commun. 1, 76 (2010). time ( exc 485 nm and em 535 nm, 10 nm bandwidth, at 25 °C). 21. Mutschler, H. & Holliger, P. 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Either 5 or 10 repeat that catalyse multiple-turnover Diels–Alder cycloadditions by using in vitro spectra were measured for buffer and bulk coacervate samples, respectively. compartmentalization. Proc. Natl Acad. Sci. USA 102, 16170–16175 (2005). Background spectra of buffer alone and bulk coacervate phase were taken under the 24. Wochner, A., Attwater, J., Coulson, A. & Holliger, P. Ribozyme-catalyzed same conditions and were subtracted from the spectra of HH-mut in buffer and transcription of an active ribozyme. Science 332, 209–212 (2011). bulk coacervate phase, respectively. All spectra were offset at 320 nm, normalised Δ θ 25. Chen, I. A., Salehi-Ashtiani, K. & Szostak, J. W. RNA catalysis in model for the cuvette pathlength and converted from A to molar ellipticity ([ ]) (deg – cm2/dmol). protocell vesicles. J. Am. Chem. Soc. 127, 13213 13219 (2005). 26. Adamala, K. P., Engelhart, A. E. & Szostak, J. W. Collaboration between primitive cell membranes and soluble catalysts. Nat. 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8 NATURE COMMUNICATIONS | (2018) 9:3643 | DOI: 10.1038/s41467-018-06072-w | www.nature.com/naturecommunications NATURE COMMUNICATIONS | DOI: 10.1038/s41467-018-06072-w ARTICLE

34. Carbonell, A., Flores, R. & Gago, S. Trans-cleaving hammerhead ribozymes Acknowledgements with tertiary stabilizing motifs: in vitro and in vivo activity against a structured Funding was provided by the Volkswagen Foundation and the MaxSynBio consortium, viroid RNA. Nucleic Acids Res. 39, 2432–2444 (2011). which is jointly funded by the Federal Ministry of Education and Research of Germany 35. Salehi-Ashtiani, K. & Szostak, J. W. In vitro evolution suggests multiple and the Max Planck Society. We thank the light microscopy facility (LMF) for their origins for the hammerhead ribozyme. Nature 414,82–84 (2001). continuous support and guidance during the FRAP experiments, and Jeff Oegema and 36. Stage-Zimmermann, T. K. & Uhlenbeck, O. C. Hammerhead ribozyme Ian Henry at the Scientific computing facility (MPI-CBG) for support with making the kinetics. RNA 4, 875–889 (1998). data available. We acknowledge the Boehringer Ingelheim Fonds for the award of a PhD 37. Osborne, E. M., Schaak, J. E. & Derose, V. J. Characterization of a native fellowship to J.M.I.-A. We thank Andre Nadler and Carl Modes for useful discussion. hammerhead ribozyme derived from schistosomes. RNA 11, 187–196 (2005). 38. Yeh, I.-C. & Hummer, G. Diffusion and electrophoretic mobility of single- Author contributions stranded rna from molecular dynamics simulations. Biophys. J. 86, 681–689 (2004). H.M., M.K., and T.-Y.D.T. conceived the project; T.-Y.D.T., H.M., B.D., J.M.I.-A., K.L.V., 39. Werner, A. Predicting translational diffusion of evolutionary conserved RNA M.Kar and V.M. undertook the experiments and data analysis; H.M., T.-Y.D.T., B.D., structures by the nucleotide number. Nucleic Acids Res. 39, e17 (2011). J.M.I.-A., K.L.V. and M.Kar discussed the results; T.-Y.D.T., H.M., B.D. and K.L.V. wrote 40. Nakano, S., Karimata, H. T., Kitagawa, Y. & Sugimoto, N. Facilitation of RNA the manuscript. enzyme activity in the molecular crowding media of cosolutes. J. Am. Chem. Soc. 131, 16881–16888 (2009). Additional information 41. Nakano, S.-I., Kitagawa, Y., Miyoshi, D. & Sugimoto, N. Hammerhead Supplementary Information accompanies this paper at https://doi.org/10.1038/s41467- ribozyme activity and oligonucleotide duplex stability in mixed solutions of 018-06072-w. water and organic compounds. FEBS Open Bio. 4, 643–650 (2014). 42. Peng, B. & Muthukumar, M. Modeling competitive substitution in a Competing interests: The authors declare no competing interests. polyelectrolyte complex. J. Chem. Phys. 143, 243133 (2015). 43. Zhang, R. & Shklovskii, B. I. Phase diagram of solution of oppositely charged Reprints and permission information is available online at http://npg.nature.com/ polyelectrolytes. Physica A 352, 216–238 (2005). reprintsandpermissions/ 44. Jia, T. Z., Hentrich, C. & Szostak, J. W. Rapid RNA exchange in aqueous two- phase system and coacervate droplets. Orig. Life Evol. Biospheres 44,1–12 (2014). Publisher's note: Springer Nature remains neutral with regard to jurisdictional claims in 45. Weinberg, M. S. & Rossi, J. J. Comparative single-turnover kinetic analyses of published maps and institutional affiliations. trans-cleaving hammerhead ribozymes with naturally derived non-conserved sequence motifs. FEBS Lett. 579, 1619–1624 (2005). 46. Han, J. & Burke, J. M. Model for general acid-base catalysis by the hammerhead ribozyme: pH-activity relationships of G8 and G12 variants at Open Access This article is licensed under a Creative Commons the putative active site. Biochemistry 44, 7864–7870 (2005). Attribution 4.0 International License, which permits use, sharing, 47. Eils, R. & Kappel, C. Fluorescence recovery after photobleaching with the adaptation, distribution and reproduction in any medium or format, as long as you give – Leica TCS SP2. Confocal Appl. Lett. 18, 1 12 (2004). appropriate credit to the original author(s) and the source, provide a link to the Creative fi 48. Leddy, H. A. & Guilak, F. Site-speci c molecular diffusion in articular cartilage Commons license, and indicate if changes were made. The images or other third party fl measured using uorescence recovery after photobleaching. Ann. Biomed. material in this article are included in the article’s Creative Commons license, unless – Eng. 31, 753 760 (2003). indicated otherwise in a credit line to the material. If material is not included in the 49. Axelrod, D., Koppel, D. E., Schlessinger, J., Elson, E. & Webb, W. W. Mobility article’s Creative Commons license and your intended use is not permitted by statutory measurement by analysis of fluorescence photobleaching recovery kinetics. – regulation or exceeds the permitted use, you will need to obtain permission directly from Biophys. J. 16, 1055 1069 (1976). the copyright holder. To view a copy of this license, visit http://creativecommons.org/ 50. Reits, E. A. J. & Neefjes, J. J. From fixed to FRAP: measuring protein mobility licenses/by/4.0/. and activity in living cells. Nat. Cell Biol. 3, E145–E147 (2001). 51. Ryabinin, V. A., Kostina, E. V. & Sinyakov, A. N. Unexpected transformation of black hole quenchers in electrophoretic purification of the fluorescein-containing © The Author(s) 2018 TaqMan probes. Nucleosides Nucleotides Nucleic Acids 36,418–427 (2017).

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