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BACKGROUND Clozapine is the most effective antipsychotic for treatment-resistant schizophrenia. Although serum clozapine levels can help guide treatment, they are underutilized owing to requirements for frequent venous blood draws and lack of immediate results. METHODS Clozapine levels measured with a novel immunoassay technology (which enables point-of-care development) were compared with those measured by standard liquid chromatography/tandem mass spectrometry (LC-MS/MS). Frozen serum aliquots of 117 samples (N = 48 patients with schizophrenia on clozapine; N = 24 patients with schizophrenia not on clozapine; N = 45 healthy controls) were sent to a national reference laboratory (NRL) for clozapine level determination by LC-MS/MS, and matching samples were subjected to novel immunoassay (3 runs). At a later date, another frozen aliquot from the same date was sent to the NRL for repeat testing. RESULTS The NRL obtained 18 false-positive clozapine results (mean 42.39 ± 32.06, range 21-159 ng/mL) in participants not on clozapine (N = 3) and healthy controls (N = 15). The immunoassay showed no false-positive clozapine results. The clozapine levels were correlated between both assays (r = 0.84, P < 0.0001), despite 16% higher clozapine levels with immunoassay (482.08 ± 270.88 ng/mL immunoassay, 414.98 ± 186.29 ng/mL LC-MS/MS [P = 0.03]). Agreement analysis using concordance correlation coefficient (CCC) for LC-MS/MS of the 2 aliquots yielded CCC = 0.869; 95% confidence interval = 0.690-0.970, whereas higher agreement results were observed for the 3 runs of immunoassay (CCC = 0.99; 95% confidence interval = 0.979-0.997). CONCLUSIONS The lack of false positives observed with immunoassay, higher repeat performance agreement, and good correlation with LC-MS/MS may indicate the more robust performance of immunoassay than that of LC-MS/MS clozapine-level determination.

Quantitative real-time PCR (qPCR) has been the standard for nucleic acid quantification as it has a large dynamic range and good sensitivity. Digital PCR is rapidly supplanting qPCR in many applications as it provides excellent quantitative precision. However, both techniques require extensive sample preparation, and highly multiplexed assays that quantify multiple targets can be difficult to design and optimize. Here we describe a new nucleic acid quantification method that we call Spatially Isolated Reactions in a Complex Array (SIRCA), a highly parallel nucleic acid preparation, amplification, and detection approach that uses superparamagnetic microbeads in an array of thousands of 100 fL microwells to simplify sample purification and reduce reagent dispensing steps. Primers, attached to superparamagnetic microbeads through a thermo-labile bond, capture and separate target sequences from the sample. The microbeads are then magnetically loaded into a microwell array such that wells predominately contain a single bead. Master mix, lacking primers, is added before sealing the reaction wells with hydrophobic oil. Thermocycling releases the primer pair from the beads during PCR amplification. At low target concentrations, most beads capture, on average, less than one target molecule, and precise, digital PCR quantification can be derived from the percentage of positive reactions. At higher concentrations, qPCR signal is used to determine the average number of target molecules per reaction, significantly extending the dynamic range beyond the digital saturation point. We demonstrate that SIRCA can quantify DNA and RNA targets using thousands of parallel reactions, achieving attomolar limits of detection and a linear dynamic range of 105. The work reported here is a first step towards multiplexed SIRCA assays.

BACKGROUND: Clozapine is an effective antipsychotic for treatment-resistant schizophrenia for which serum levels could guide therapy decisions. It is underutilized because it requires venous draws and several-day determination time with high performance liquid chromatography-tandem mass spectrometry (LC/MS-MS). This project evaluates if clozapine measured with a novel immunoassay technology, which could be developed into a fingerstick test, correlate with LC/MS-MS. This project also assesses the impact of demographic and clinical variables on assay results. METHODS: 117 serum samples (N=48 with schizophrenia on clozapine, N=24 with schizophrenia not on clozapine and N=45 healthy controls) were included. One aliquot was sent to a national reference laboratory (NRL) for determination by LC/MS-MS and another sent to Saladax for immunoassay (MyCare® Psychiatry Clozapine Assay Kit). The agreement was compared using Concordance Correlation Coefficient (CCC). Participants’ age, sex, race-ethnicity, smoking status, ascorbic acid, co-medications and complete metabolic panel were collected. Linear regression and mixed model were performed for both technologies correspondingly to examine the impact of these variables. RESULTS: Clozapine levels by the NRL had 19 false positive readings (mean 48.8 ± 14.3, range 21–138 ng/ml) in schizophrenia participants not on clozapine (N=3) and healthy controls (N=16). The immunoassay had no false positive results. A mixed effects model (with a time average of the NRL and 3 immunoassay runs), yielded a strong Pearson correlation (r=0.843, p

SHAPE chemistries exploit small electrophilic reagents that react with the 2′-hydroxyl group to interrogate RNA structure at single-nucleotide resolution. Mutational profiling (MaP) identifies modified residues based on the ability of reverse transcriptase to misread a SHAPE-modified nucleotide and then counting the resulting mutations by massively parallel sequencing. The SHAPE-MaP approach measures the structure of large and transcriptome-wide systems as accurately as for simple model RNAs. This protocol describes the experimental steps, implemented over three days, required to perform SHAPE probing and construct multiplexed SHAPE-MaP libraries suitable for deep sequencing. These steps include RNA folding and SHAPE structure probing, mutational profiling by reverse transcription, library construction, and sequencing. Automated processing of MaP sequencing data is accomplished using two software packages. ShapeMapper converts raw sequencing files into mutational profiles, creates SHAPE reactivity plots, and provides useful troubleshooting information, often within an hour. SuperFold uses these data to model RNA secondary structures, identify regions with well-defined structures, and visualize probable and alternative helices, often in under a day. We illustrate these algorithms with the E. coli thiamine pyrophosphate riboswitch, E. coli 16S rRNA, and HIV-1 genomic RNAs. SHAPE-MaP can be used to make nucleotide-resolution biophysical measurements of individual RNA motifs, rare components of complex RNA ensembles, and entire transcriptomes. The straightforward MaP strategy greatly expands the number, length, and complexity of analyzable RNA structures.

Currently, reliable valving on integrated microfluidic devices fabricated from rigid materials is confined to expensive and complex methods. Freeze-thaw valves (FTVs) can provide a low cost, low complexity valving mechanism, but reliable implementation of them has been greatly hindered by the lack of ice nucleation sites within the valve body's small volume. Work to date has required very low temperatures (on the order of -40 °C or colder) to induce freezing without nucleation sites, making FTVs impractical due to instrument engineering challenges. Here, we report the use of ice-nucleating proteins (INPs) to induce ice formation at relatively warm temperatures in microfluidic devices. Microfluidic channels were filled with buffers containing femtomolar INP concentrations from Pseudomonas syringae. The channels were cooled externally with simple, small-footprint Peltier thermoelectric coolers (TECs), and the times required for channel freezing (valve closure) and thawing (valve opening) were measured. Under optimized conditions in plastic chips, INPs made sub-10 s actuations possible at TEC temperatures as warm as -13 °C. Additionally, INPs were found to have no discernible inhibitory effects in model enzyme-linked immunosorbent assays or polymerase chain reactions, indicating their compatibility with microfluidic systems that incorporate these widely used bioassays. FTVs with INPs provide a much needed reliable valving scheme for rigid plastic devices with low complexity, low cost, and no moving parts on the device or instrument. The reduction in freeze time, accessible actuation temperatures, chemical compatibility, and low complexity make the implementation of compact INP-based FTV arrays practical and attractive for the control of integrated biochemical assays.

Many biological processes are RNA-mediated, but higher-order structures for most RNAs are unknown, which makes it difficult to understand how RNA structure governs function. Here we describe selective 2'-hydroxyl acylation analyzed by primer extension and mutational profiling (SHAPE-MaP) that makes possible de novo and large-scale identification of RNA functional motifs. Sites of 2'-hydroxyl acylation by SHAPE are encoded as noncomplementary nucleotides during cDNA synthesis, as measured by massively parallel sequencing. SHAPE-MaP-guided modeling identified greater than 90% of accepted base pairs in complex RNAs of known structure, and we used it to define a new model for the HIV-1 RNA genome. The HIV-1 model contains all known structured motifs and previously unknown elements, including experimentally validated pseudoknots. SHAPE-MaP yields accurate and high-resolution secondary-structure models, enables analysis of low-abundance RNAs, disentangles sequence polymorphisms in single experiments and will ultimately democratize RNA-structure analysis.

Structured RNA elements within messenger RNA often direct or modulate the cellular production of active proteins. As reviewed here, RNA structures have been discovered that govern nearly every step in protein production: mRNA production and stability; translation initiation, elongation, and termination; protein folding; and cellular localization. Regulatory RNA elements are common within RNAs from every domain of life. This growing body of RNA-mediated mechanisms continues to reveal new ways in which mRNA structure regulates translation. We integrate examples from several different classes of RNA structure-mediated regulation to present a global perspective that suggests that the secondary and tertiary structure of RNA ultimately constitutes an additional level of the genetic code that both guides and regulates protein biosynthesis.

Secondary structure plays critical roles in nucleic acid function. Mismatches in DNA can lead to mutation and disease, and some mismatches involve a protonated base. Among protonated mismatches, A(+).C wobble pairs form near physiological pH and have relatively minor effects on helix geometry, making them especially important in biology. Herein, we investigate effects of helix position, temperature, and ionic strength on pK(a) shifting in A(+).C wobble pairs in DNA. We observe that pK(a) shifting is favored by internal A(+).C wobbles, which have low cooperativities of folding and make large contributions to stability, and disfavored by external A(+).C wobbles, which have high folding cooperativities but make small contributions to stability. An inverse relationship between pK(a) shifting and temperature is also found, which supports a model in which protonation is enthalpically favored overall and entropically correlated with cooperativity of folding. We also observe greater pK(a) shifts as the ionic strength decreases, consistent with anticooperativity between proton binding and counterion-condensed monovalent cation. Under the most favorable temperature and ionic strength conditions tested, a pK(a) of 8.0 is observed for the A(+).C wobble pair, which represents an especially large shift ( approximately 4.5 pK(a) units) from the unperturbed pK(a) value of adenosine. This study suggests that protonated A(+).C wobble pairs exist in DNA under biologically relevant conditions, where they can drive conformational changes and affect replication and transcription fidelity.

The hepatitis delta virus (HDV) ribozyme uses the nucleobase C75 and a hydrated Mg(2+) ion as the general acid-base catalysts in phosphodiester bond cleavage at physiological salt. A mechanistic framework has been advanced that involves one Mg(2+)-independent and two Mg(2+)-dependent channels. The rate-pH profile for wild-type (WT) ribozyme in the Mg(2+)-free channel is inverted relative to the fully Mg(2+)-dependent channel, with each having a near-neutral pKa. Inversion of the rate-pH profile was used as the crux of a mechanistic argument that C75 serves as general acid both in the presence and absence of Mg(2+). However, subsequent studies on a double mutant (DM) ribozyme suggested that the pKa observed for WT in the absence of Mg(2+) arises from ionization of C41, a structural nucleobase. To investigate this further, we acquired rate-pH/pD profiles and proton inventories for WT and DM in the absence of Mg(2+). Corrections were made for effects of ionic strength on hydrogen ion activity and pH meter readings. Results are accommodated by a model wherein the Mg(2+)-free pKa observed for WT arises from ionization of C75, and DM reactivity is compromised by protonation of C41. The Brønsted base appears to be water or hydroxide ion depending on pH. The observed pKa's are related to salt-dependent pH titrations of a model oligonucleotide, as well as electrostatic calculations, which support the local environment for C75 in the absence of Mg(2+) being similar to that in the presence of Mg(2+) and impervious to bulk ions. Accordingly, the catalytic role of C75 as the general acid does not appear to depend on divalent ions or the identity of the Brønsted base.

Secondary structural motifs play essential roles in the folding and function of RNA and DNA molecules. Previous work from our lab compared the folding of small DNA and RNA hairpin loops containing a sheared GA pair [Moody, E. M., Feerar, J. C., and Bevilacqua, P. C. (2004) Biochemistry 43, 7992-7998]. We found that the small DNA hairpins fold in a highly cooperative manner with indirect coupling, while their RNA counterparts fold in a much less cooperative fashion and display direct coupling. Herein, we extend this study to the double-stranded helix. We carried out double mutant cycles on base pairs having identical nearest-neighbor contexts but located in either external or internal helical registers. In the external register, both RNA and DNA exhibit extensive folding cooperativity between the penultimate and terminal base pair, which is independent of mismatch identity. In contrast, DNA exhibits virtually no folding cooperativity in the center of the helix, while RNA maintains substantial coupling, which is dependent on mismatch identity. Two models account for these non-nearest-neighbor effects: one involves the unfavorable entropy of helix initiation common to DNA and RNA, and the other involves steric and electrostatic strain peculiar to RNA. These data show that RNA can display cooperativity less than, greater than, or equal to that of DNA depending on context and position.

RNA plays essential roles in much of biology. These functions are dictated by structures mediated by hydrogen bonding, stacking, electrostatics, and steric interactions. Roles of unsatisfied hydrogen bond functionalities in these structures are less well understood. Herein, we evaluated the energetic contributions of unsatisfied hydrogen bonding groups by placing chemically modified substituents in select internal positions in RNA helices and conducting thermodynamic studies. We find that unsatisfied carbonyl groups make exceptional contributions to structure formation (approximately 3 kcal/mol in free energy), most likely due to a combination of strain and dehydration effects. Thus, unsatisfied hydrogen bonding groups are likely key determinants in the folding energetics and specificity of many RNA and DNA molecules and may be especially important in tertiary structure interactions.

Small ribozymes such as the hairpin, hammerhead, VS, glm S, and hepatitis delta virus (HDV) are self-cleaving RNAs that are typically characterized by kinetics and structural methods. Working with these RNAs requires attention to numerous experimental details. In this chapter we focus on four different experimental aspects of ribozyme studies: preparing the RNA, mapping its structure with reverse transcription and end-labeled techniques, solvent isotope experiments, and co-transcriptional cleavage assays. Although the focus of these methods is the HDV ribozyme, the methods should be applicable to other ribozymes.

Small ribozymes such as the hairpin, hammerhead, VS, glm S, and hepatitis delta virus (HDV) are self-cleaving RNAs that are typically characterized by kinetics and structural methods. Working with these RNAs requires attention to numerous experimental details. In this chapter we focus on four different experimental aspects of ribozyme studies: preparing the RNA, mapping its structure with reverse transcription and end-labeled techniques, solvent isotope experiments, and co-transcriptional cleavage assays. Although the focus of these methods is the HDV ribozyme, the methods should be applicable to other ribozymes.

RNA SHAPE chemistry yields quantitative, single nucleotide resolution structural information based on the reaction of the 2′-hydroxyl group of conformationally flexible nucleotides with electrophilic SHAPE reagents. However, SHAPE technology has been limited by the requirement that sites of RNA modification be detected by primer extension. Primer extension results in loss of information at both the 5′ and 3′ ends of an RNA and requires multiple experimental steps. Here we describe RNase-detected SHAPE (Selective 2′-Hydroxyl Acylation analyzed by Protection from Exoribonuclease) that uses a processive, 3′→5′ exoribonuclease, RNase R, to detect covalent adducts in 5′-end labeled RNA in a one-tube experiment. RNase R degrades RNA but stops quantitatively three and four nucleotides 3′ of a nucleotide containing a covalent adduct at the ribose 2′-hydroxyl or the pairing face of a nucleobase, respectively. We illustrate this technology by characterizing ligand-induced folding for the E. coli thiamine pyrophosphate riboswitch RNA. RNase-detected SHAPE is a facile, two-day approach that can be used to analyze diverse covalent adducts in any RNA molecule, including short RNAs not amenable to analysis by primer extension and RNAs with functionally important structures at their 5′ or 3′ ends.

Double and triple mutant thermodynamic cycles provide a means to dissect the cooperativity of RNA and DNA folding at both the secondary and tertiary structural levels through use of the thermodynamic box or cube. In this article, we describe three steps for applying thermodynamic cycles to nucleic acid folding, with considerations of both conceptual and experimental features. The first step is design of an appropriate system and development of hypotheses regarding which residues might interact. Next is implementing this design in terms of a tractable experimental strategy, with an emphasis on UV melting. The final step, and the one we emphasize the most, is interpreting mutant cycles in terms of coupling between specific residues in the RNA or DNA. Coupling free energy in the absence and presence of changes elsewhere in the molecule is discussed in terms of specific folding models, including stepwise folding and concerted changes. Last, we provide a practical section on the use of commercially available software (KaleidaGraph) to fit melting data, along with a consideration of error propagation. Along the way, specific examples are chosen from the literature to illustrate the methods. This article is intended to be accessible to the biochemist or biologist without extensive thermodynamics background.

Double and triple mutant thermodynamic cycles provide a means to dissect the cooperativity of RNA and DNA folding at both the secondary and tertiary structural levels through use of the thermodynamic box or cube. In this article, we describe three steps for applying thermodynamic cycles to nucleic acid folding, with considerations of both conceptual and experimental features. The first step is design of an appropriate system and development of hypotheses regarding which residues might interact. Next is implementing this design in terms of a tractable experimental strategy, with an emphasis on UV melting. The final step, and the one we emphasize the most, is interpreting mutant cycles in terms of coupling between specific residues in the RNA or DNA. Coupling free energy in the absence and presence of changes elsewhere in the molecule is discussed in terms of specific folding models, including stepwise folding and concerted changes. Last, we provide a practical section on the use of commercially available software (KaleidaGraph) to fit melting data, along with a consideration of error propagation. Along the way, specific examples are chosen from the literature to illustrate the methods. This article is intended to be accessible to the biochemist or biologist without extensive thermodynamics background.

The RNA World hypothesis posits that life emerged from self-replicating RNA molecules. For any single biopolymer to be the basis for life, it must both store information and perform diverse functions. It is well known that RNA can store information. Advances in recent years have revealed that RNA can exhibit remarkable functional versatility as well. In an effort to judge the functional versatility of RNA and thereby the plausibility that RNA was at one point the basis for life, a statistical chemical approach is adopted. Essential biological functions are reduced to simple molecular models in a minimalist, biopolymer-independent fashion. The models dictate requisite states, populations of states, and physical and chemical changes occurring between the states. Next, equations are derived from the models, which lead to complex phenomenological constants such as observed and functional constants that are defined in terms of familiar elementary chemical descriptors: intrinsic rate constants, microscopic ligand equilibrium constants, secondary structure stability, and ligand concentration. Using these equations, simulations of functional behavior are performed. These functional models provide practical frameworks for fitting and organizing real data on functional RNAs such as ribozymes and riboswitches. At the same time, the models allow the suitability of RNA as a basis for life to be judged. We conclude that RNA, while inferior to extant proteins in most, but not all, functional respects, may be more versatile than proteins, performing a wider range of elementary biological functions at a tolerable level. Inspection of the functional models and various RNA structures uncovers several surprising ways in which the nucleobases can conspire to afford chemical catalysis and evolvability. These models support the plausibility that RNA, or a closely related informational biopolymer, could serve as the basis for a fairly simple form of life.

Authors: Kady-Ann Steen, Nathan A. Siegfried & Kevin M. Weeks ### Abstract SHAPE (selective 2’-hydroxyl acylation analyzed by primer extension) chemistry is a powerful approach for single-nucleotide resolution analysis of RNA secondary and tertiary structure. SHAPE uses hydroxyl-selective electrophiles to react with and form covalent adducts at the ribose 2’-hydroxyl group. The most useful and robust reagent for routine SHAPE experiments is 1-methyl-7-nitroisatoic anhydride (1M7). This protocol describes a straightforward synthesis of 1M7 from commercially available precursors. The synthesis of 1-methyl-7-nitroisatoic anhydride was originally described by Mortimer et al. . The reaction scheme is:   *Amounts of each starting material can be varied using the above Table*. ### Reagents 1. 4-nitroisatoic anhydride (4NIA) (AstaTech, cat. no. 69441) - Sodium hydride (NaH) – 60% dispersion in mineral oil (Sigma-Aldrich, cat. no. 452912) - Methyl iodide (MeI) (Sigma-Aldrich, cat. no. 289566) - Anhydrous dimethylfluoride (DMF) (Sigma-Aldrich, cat. no. 227056) - HCl (Fisher Scientific, cat. no. A144) - H2O - Ether (Fisher Scientific, cat. no. E138) **REAGENT SETUP** - H2O and HCl should be ice cold before use. ### Equipment 1. Round bottom flasks (250 and 500 mL) - Bunsen burner - N2 tank and gas line - Teflon-coated stirring bar - Rubber septa - Disposable syringes (1 mL, 20 mL, 60 mL) - Disposable needles (18 gauge) - Stirring plate - Buchner funnel - Vacuum rotary evaporator - Watch glass - Oven maintained at 90 °C **EQUIPMENT SETUP** - All glassware used in the described reactions are washed with soap and water, rinsed with water, rinsed with acetone, and flame-dried over Bunsen burner. Rubber septa are immediately placed over flasks, flasks are flushed with N2, and allowed to cool. ### Procedure 1. Dissolve 4NIA in 60 mL DMF in a 250 mL flame-dried round bottom flask under N2. - In a separate 500 mL flame-dried round bottom flask under N2, make a slurry of NaH in 15 mL DMF and stir. - Slowly add 4NIA solution dropwise to slurry of NaH in DMF. Stir for 5 minutes. - Slowly add MeI dropwise and then stir at room temperature for 4 hours. - Pour reaction into 100 mL *ice cold* 1N HCl. - Filter resulting bright orange precipitate by vacuum filtration in a Buchner funnel. - Rinse precipitate twice with *ice cold* water, followed by 3 rinses with ether. - Dry overnight in oven on watch glass. ### Anticipated Results **Analytical data** Yield ~80%; bright orange powdery solid. 1H NMR [400 MHz, CO(CD3)2]: σ 3.69 (s, 3H, -NCH3-), 8.12 (dd, J=8.8 Hz, 2 Hz, 1 H, ArH), 8.2 (d, J=2hz, 1h, ArH), 8.34 (d, J=8.4 Hz, 1 H, ArH). ### Associated Publications **Selective 2′-hydroxyl acylation analyzed by protection from exoribonuclease (RNase-detected SHAPE) for direct analysis of covalent adducts and of nucleotide flexibility in RNA**. Kady-Ann Steen, Nathan A Siegfried, and Kevin M Weeks. *Nature Protocols* 6 (11) 1683 - 1694 [doi:10.1038/nprot.2011.373](http://dx.doi.org/10.1038/nprot.2011.373) ### Author information **Kady-Ann Steen, Nathan A. Siegfried & Kevin M. Weeks**, Department of Chemistry, University of North Carolina, Chapel Hill, NC 27599-3290 Correspondence to: Kevin M. Weeks (weeks@unc.edu) *Source: [Protocol Exchange](http://www.nature.com/protocolexchange/protocols/2177) (2011) doi:10.1038/protex.2011.255. Originally published online 3 November 2011*.