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. Author manuscript; available in PMC: 2023 Apr 1.
Published in final edited form as: Biomol NMR Assign. 2022 Mar 12;16(1):153–158. doi: 10.1007/s12104-022-10073-8

Backbone and sidechain 1H, 15N and 13C resonance assignments of the free and RNA-bound tandem zinc finger domain of the tristetraprolin family member from Selaginella moellendorffii

Stephanie N Hicks 1, Ronald A Venters 2, Perry J Blackshear 1,3,4
PMCID: PMC9196822  NIHMSID: NIHMS1804422  PMID: 35279790

Abstract

Members of the tristetraprolin (TTP) family of RNA binding proteins (RBPs) regulate the metabolism of a variety of mRNA targets. In mammals, these proteins modulate many physiological processes, including immune cell activation, hematopoiesis, and embryonic development. Regulation of mRNA stability by these proteins requires that the tandem zinc finger (TZF) domain binds initially and directly to target mRNAs, ultimately leading to their deadenylation and decay. Proteins of this type throughout eukarya possess a highly conserved TZF domain, suggesting that they are all capable of high-affinity RNA binding. However, the mechanism of TTP-mediated mRNA decay is largely undefined. Given the vital role that these TTP family proteins play in maintaining RNA homeostasis throughout eukaryotes, we focused here on the first, key step in this process: recognition and binding of the TZF domain to target RNA. For these studies, we chose a primitive plant, the spikemoss Selaginella moellendorffii, which last shared a common ancestor with humans more than a billion years ago. Here we report the near complete backbone and side chain resonance assignments of the spikemoss TZF domain, including: 1) the assignment of the RNA-TZF domain complex, representing one of only two data sets currently available for the entire TTP family of proteins; and 2) the first NMR resonance assignments of the entire TZF domain, in the RNA-free form. This work will serve as the basis for further NMR structural investigations aimed at gaining insights into the process of RNA recognition and the mechanisms of TTP-mediated mRNA decay.

Keywords: RNA binding proteins, tristetraprolin, zinc finger, Selaginella moellendorffii, spikemoss, RNA decay

Biological context

RNA binding proteins (RBPs) interact with RNA through a variety of domains. Zinc finger (ZF)-containing RBPs possess cysteine and histidine residues, in various arrangements, to coordinate zinc, a requirement for their stability and, thus, interaction with RNA. The tristetraprolin (TTP) family of RBPs contains a tandem zinc finger (TZF) domain that is defined by two highly conserved cysteine and histidine motifs, with the arrangement CX8CX5CX3H, that interact directly with target mRNAs to initiate mRNA metabolism (Lai et al. 2000). Experiments in mice have shown that mammalian TTP family members are vital to maintaining the cellular balance of transcripts encoding proteins involved in processes as diverse as innate immunity, formation of the umbilical circulation, definitive hematopoiesis, ovarian development and female fertility, and placental physiology (Taylor et al. 1996; Stumpo et al. 2004; Stumpo et al. 2009; Stumpo et al. 2016). Non-vertebrate members of this protein family appear to control important physiological processes, such as iron metabolism and cell-cell interactions, using similar mechanisms in the yeasts S. cerevisiae, S. pombe, and C. albicans, the fruit fly D. melanogaster, and the social ameba D. discoideum (Puig et al. 2005; Wells et al. 2012; Cuthbertson et al. 2008; Wells et al. 2015; Choi et al. 2014; Bai et al. 2021). Due to the importance of members of the TTP family of proteins in regulating RNA homeostasis, and other vital cellular processes, it is imperative to understand the process of the initial RNA recognition event, and the mechanism of mRNA decay mediated by this family of RNA binding proteins.

To mediate mRNA turnover, TTP family members must first bind to AU-rich elements (AREs) within the 3’-untranslated regions (UTR) of messenger RNA (mRNA) targets. This high affinity binding is the prerequisite for the next steps, i.e., the recruitment of enzymes and other proteins that can catalyze removal of the polyA tail by 3’−5’ exonuclease activities, a process known as deadenylation. Although specific mRNA targets have yet to be elucidated for many of the TTP family members, several cytokine and chemokine transcripts in the mouse have been identified as direct targets of TTP itself, including the transcripts encoding tumor necrosis factor (TNF), granulocyte-macrophage colony-stimulating factor (GM-CSF), C-X-C motif chemokine ligand 1 (CXCL1), C-X-C motif chemokine ligand 2 (CXCL2), immediate early response 3 (IER3), and others (Lai et al. 1999; Carballo et al. 2000). Although it is clear that TTP-mediated mRNA decay is exceedingly important for regulation of inflammation in mammals, the mechanisms by which it stimulates mRNA deadenylation and decay remain largely unclear.

The TZF domain is an essential 64 amino acid RNA recognition element within the TTP family of proteins and is highly conserved, with approximately 65% amino acid identity between organisms whose last common ancestors lived over 1 billion years ago (Lai et al. 2019a). In contrast, there is generally little conservation among other eukaryotic family members in other regions of these proteins, with the exception of a short sequence at the extreme C-terminus in some species that is involved in interactions with the CCR4-NOT complex (Fabian et al. 2013; Lai et al. 2019b). Importantly, the TZF domain alone has been shown to be necessary and sufficient for direct binding to RNA, with the optimal binding sequence being identified as 5’-UUAUUUAUU-3’ in the few organisms in which it has been studied. It is clear that many, if not most, members of this protein family are vital for regulating mRNA processing and translation in cells. However, the vast majority of TTP family proteins have not been studied in detail, leaving many unanswered questions, including: the molecular mechanisms of binding to mRNA targets, particularly the interactions between the individual zinc fingers and RNA, and the interactions between the zinc fingers; the equilibrium of these interactions; the target sequence specificity in evolutionarily widely separated organisms; the effects of sequence “drift” within the TZF domain, as has been observed in some groups of species; and the potential identities and functions of interacting protein partners that might modulate the primary RNA binding activity. To gain further insights into the mechanisms of TTP family protein-mediated mRNA decay, and to begin to address some of the questions pertaining to RNA recognition and binding to the TZF domain, we report here the complete NMR resonance assignments of the TZF domain from a TTP family member, both in the RNA bound and unbound states.

For the current study, we used the TZF domain from the TTP family member expressed in the primitive spikemoss Selaginella moellendorffii (S. moellendorffii), commonly known as Gemmiferous spikemoss. This organism was selected in part because of its extreme evolutionary distance from humans, estimated at greater than 1.5 billion years since the last common ancestor; the exceptionally small size of the intact protein, since at 119 AA it is the shortest TTP family member identified to date; the presence of a potential CNOT1 interacting domain at its very C-terminus; and its solubility and stability during bacterial expression and purification, unusual for this family of proteins. The isolated TZF domain (residues 10 – 76 from the full-length protein; 67 residues) from the spikemoss protein was used for the solution NMR studies reported here, and has allowed, for the first time in members of this family, residue assignments for the RNA-free form of the entire TZF domain. Resonance assignments are also presented for the RNA-bound form of the spikemoss TZF domain, and compared to the assignments for the human TIS11d-RNA (BioMagResBank; http://www.bmrb.wisc.edu; accession code 6005) complex, the only available assignments for a TTP family member TZF domain-RNA complex for this family of proteins prior to this study (Hudson et al. 2004). The work described here will not only serve as the basis for our initial structural characterizations of the RNA-bound and free TZF domains in solution, but also the initial exploration into the process of RNA recognition, and the role that protein dynamics plays in this process. These data should provide new insights into the mechanisms of mRNA decay promoted by this family of RNA binding proteins.

Methods and Experiments

Plasmid constructs

DNA for the full-length TTP family member from S. moellendorffii (RefSeq accession number XP_002980683.1) was synthesized by Genewiz (South Plainfield, NJ). DNA encoding the TZF domain of S. moellendorffii, corresponding to amino acids 10 – 76 of the full-length protein, was PCR amplified and cloned, using the SspI site, into a modified pET28/30 vector, encoding an N-terminal six residue histidine tag immediately followed by a maltose binding protein (MBP) tag and a tobacco etch viral (TEV) protease cleavage site, as described previously (Wells et al. 2015). Fidelity of the expression plasmid was confirmed by DNA sequencing.

Expression and purification of labeled proteins

After transformation of the plasmid encoding the TZF domain of S. moellendorffii into E. coli BL21 Star (DE3) cells (Invitrogen, Waltham, Massachusetts), the cells were plated onto LB agar containing 100 μg/mL ampicillin. Bacterial cultures were grown at 37°C in modified M9 minimal medium containing 1 g/L 15NH4Cl and 2 g/L 13C6-D-glucose (Cambridge Isotopes, Tewksbury, MA) and 100 μg/mL ampicillin. When the OD600 reached 0.6, 1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG) was used to induce protein expression at 37°C for 4 h. Cells were centrifuged at 3600 rpm at 4°C for 15 min and pellets were stored at −20°C.

Pelleted cells were resuspended in 50 mL of buffer containing 20 mM sodium phosphate, pH 8.0, 100 mM potassium chloride, 10 mM imidazole, 5% glycerol, and one EDTA-free protease inhibitor tablet (Roche, Basel, Switzerland) and lysed by sonication for 30 s intervals, for a total of 4 intervals, on ice. Following centrifugation at 35,000 rpm at 4°C for 30 min, supernatants were applied to a 5 mL HisTrap HP column (GE Healthcare), washed with 5 column volumes of buffer and eluted in the same buffer containing 400 mM imidazole. After incubation with TEV protease overnight at 4°C in the presence of 3 mM dithiothreitol, proteins were diluted into buffer A containing 20 mM sodium phosphate, pH 7.6, 30 mM potassium chloride, 25 μM zinc sulfate, 2 mM 2-mercaptoethanol, and 5% glycerol. Subsequently, protein was loaded onto a HiTrap Heparin HP 5 mL column (GE Healthcare) and eluted in a linear gradient from 0% to 80% buffer B (buffer A + 1M KCl). After concentration utilizing a 3,000 MWCO Vivaspin 20 centrifugal filtering device (GE Healthcare), protein was applied to a Superdex 200 10/300 GL size-exclusion column (GE Healthcare) equilibrated in 20 mM sodium phosphate, pH 7.6, 100 mM potassium chloride, 25 μM zinc sulfate, 2 mM 2-mercaptoethanol, and 5% glycerol. Fractions containing purified TZF domain were pooled and concentrated using a 3,000 MWCO Vivaspin 20 centrifugal filtering device. Purified protein was buffer exchanged into 20 mM sodium phosphate, pH 6.2, 100 mM sodium chloride, 25 μM zinc sulfate, 2 mM 2-mercaptoethanol, and 5% glycerol. Concentrated proteins were stored at −80°C after flash freezing in liquid nitrogen.

Preparation of TZF-RNA complex

The RNA 9-mer, 5’-UUAUUUAUU-3’, was purchased from GE Dharmacon (Lafayette, CO), and was deprotected in the supplied buffer and vacuum dried. After resuspension into buffer containing 20 mM sodium phosphate, pH 6.2, 100 mM potassium chloride, 25 μM zinc sulfate, 2 mM 2-mercaptoethanol, and 5% glycerol, RNA was mixed with uniformly labeled [13C, 15N] TZF domain in a 1:1.2 molar ratio.

NMR spectroscopy

[13C, 15N] uniformly labeled spikemoss TZF domain samples at concentrations between 0.4 and 0.6 mM, containing 95% H2O/5% D2O, were prepared and transferred to 5 mm Shigemi tubes (Shigemi, Inc., Allison Park, USA) for data collection at 25°C, as calibrated using a standard methanol sample. Assignment data for both the apo TZF domain and the RNA-TZF domain complex were collected on a Bruker Avance 14.1T (600MHz) spectrometer equipped with a triple-resonance cryoprobe and pulsed-field Z-gradient. Backbone resonance assignment data were collected using a set of sparse sampling NMR experiments, including HNCO, HN(CA)CO, HNCA, HN(CO)CA, HN(CA)CB, HN(COCA)CB, HA(CA)NH, and HA(CACO)NH (Venters et al. 2005; Coggins and Zhou 2008). Sidechain resonance assignments for the apo and RNA-bound form of the TZF domain were determined using a non-uniform sampling 4D HCC(CO)NH experiment and, after exchange into buffer containing 100% D2O, a non-uniform sampling 4D HCCH TOCSY experiment (Coggins and Zhou 2008). When comparing the non-uniform sampled data sets with conventional data collected at the same overall resolution, the percent sampling used herein were 5.4% for all of the 3D experiments and 0.8% for all of the 4D experiments. Data were processed using NMRPipe (Delaglio et al. 1995). NMRViewJ version 8.0 (Johnson and Blevins 1994) was used for peak picking and assignment, after referencing to an external 2,2-dimethyl-2-silapentane-5-sulfonate (DSS) standard. Initial resonance assignments were determined using PINE (Lee and Markley 2018) and AutoAssign (Zimmerman et al. 1997), with discrepancies and missing spins assigned manually. The backbone and sidechain assignments of the TZF domain-RNA complex were determined de novo and are completely independent of the RNA-free TZF domain assignments.

Extent of assignments and data deposition

Both the RNA-free and RNA-bound forms of the TZF domain from the S. moellendorffii TTP protein (residues 10 – 76; 8.059 Da) gave rise to well dispersed 1H-15N HSQC spectra (Figs. 1a and b, respectively). The resonance assignments for the three additional amino acid residues at the N-terminus, present because of the TEV site for cleavage of the affinity tag used for purification, are not presented here. The final levels of completeness obtained for the backbone resonance assignments of both the RNA-free and RNA-bound proteins were 97% for the non-proline backbone 1HN and 15N resonances, and 100% for the 13CO, 13Cα, 13Cβ, 1Hα, and 1Hβ nuclei. In both forms of the protein, it was not possible to assign the backbone 1HN and 15N resonances for Asn29 and Gly36 residues, presumably because of intermediate exchange dynamics. All of the expected sidechain 1HC and 13C resonances were assigned for the RNA-free TZF domain, including the aromatic ring resonances, with the exception of Trp19 and Lys50 Cδ. Incomplete sidechain and aromatic resonance assignments in the RNA-bound form of the protein were realized for Trp19, Phe33, His46 Cδ, Lys48, Arg68 Cδ and Phe71 Cδ. All of the Asn Nδ and Gln Nε sidechain 1HN and 15N resonances were assigned in both forms of the protein. In the RNA-bound form of this TZF domain, six of the possible seven Arg Nε resonances are seen folded into the HSQC spectrum (Fig. 1b) and have been assigned. In contrast, the Arg Nε resonances are not observed in the RNA-free form of the protein, presumably due to exchange broadening with the solvent. Aromatic 1HC and 13C and sidechain 1HN and 15N resonances were assigned using non-uniformly sampled 4D 13C/13C and 13C/15N NOE data sets. Backbone 1H-15N assignments are indicated in the 2D 1H-15N HSQC spectra (Fig. 1). Both fingers, ZF1 and ZF2, comprising the TZF domain of the spikemoss protein, were observed and assigned in solution. Prior to this study, only two partial assignments of the RNA-free form of the TZF domain have been reported for the entire TTP family of proteins. Both prior reports only successfully assigned the first finger, ZF1, of human TTP (Blackshear et al. 2003) and mouse TTP (Nup475/TIS11) (BioMagResBank; http://www.bmrb.wisc.edu; accession code 5525) (Amann et al. 2003). Comparison of the RNA-free and RNA-bound forms of the TZF domain of the spikemoss protein reveals that there are significant chemical shift perturbations within both ZF domains, upon RNA (5’-UUAUUUAUU-3’; 2739 Da) binding. Those residues with chemical shift perturbations at least one standard deviation above the overall average are highlighted in Figure 1.

Fig. 1. 1H-15N HSQC spectra of the spikemoss a) apo TZF domain and the b) TZF-RNA complex.

Fig. 1

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Assignments for the RNA-free (black) and the RNA-bound (blue) TZF domain are numbered according to their context within the full-length protein. Selected peaks with chemical shift perturbations at least one standard deviation larger than the average, upon RNA binding, are labeled in red. c) Amide chemical shift perturbation of the TZF domain upon binding RNA. Residues with 1HN-15N chemical shift perturbations of at least one standard deviation above the mean are highlighted in red.

The spikemoss TTP family protein TZF domain studied here shares 65 % amino acid identity with the previously reported human TTP family member, TIS11d (ZFP36L2) (BioMagResBank; http://www.bmrb.wisc.edu; accession code 6005) (Hudson et al. 2004). Comparison of the assignments of the TZF-RNA complexes reveal that both TTP family members have well-dispersed HSQC spectra in the presence of RNA, and that near complete assignment of both zinc finger domains is possible in the presence of RNA. In addition, equivalent residues in the two complexes are seen to experience significant chemical shift perturbation upon RNA binding. The secondary structure propensity predicted by TALOS+, based on backbone 1H, 15N, 13C, Cα, and Cβ chemical shifts of the spikemoss TZF domain, indicates α-helical (R17 - T22 in ZF1 and T56 - A59 in ZF2) and 310-helical regions (Y27 - N29 in ZF1, K37 - D39 at the interface between ZF1 and the linker, and Y65 - K67 in ZF2) equivalent to those observed in the NMR structure of TIS11d-RNA complex. Furthermore, the linker region between the two fingers in the TZF domain of spikemoss is unstructured (residues D39 - E52), in both the presence and absence of RNA, similar to the linker region in the TIS11d-RNA complex (Fig. 2).

Fig. 2. Sequence and secondary structure of the TZF domain of the tristetraprolin (TTP) family member from spikemoss.

Fig. 2

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a) Sequence of the purified TZF domain used for data collection, numbered based on the position within the full-length protein. Three additional residues, serine, asparagine, and alanine, are added to the N-terminus of the TZF domain, due to a TEV cleavage site that was introduced for removal of a tandem 6 residue histidine and MBP tag for affinity purification. However, these amino acids are not included in the assignments. Secondary structure propensity of the TZF domain of the spikemoss protein in the b) RNA-free and c) RNA-bound state, as predicted by TALOS+, based on backbone 1H, 15N, 13C, Cα, and Cβ chemical shifts. Propensities to form alpha helix (blue bars) or beta sheets (red bars), are plotted versus residue number.

The 1H, 15N, and 13C resonance assignments for the backbone and sidechain atoms of the RNA-free and RNA-bound form of the TZF domain of the spikemoss TTP protein have been deposited in the BioMagResBank database (http://www.bmrb.wisc.edu) under accession codes 51209 and 51210.

Conclusion

We present our assignments of the RNA-free and RNA-bound forms of the TZF domain of the TTP family member from the spikemoss S. moellendorffii. These studies represent the first near-complete assignment of the entire TZF domain, in an RNA-free state, of a TTP family member. We also compare the RNA-bound TZF domain of spikemoss protein to that of the only other TZF domain-RNA complex assignment currently available for the TTP family, from human TIS11d (ZFP36L2). This work will be used as the basis for: 1) further structural work on the RNA-free and RNA-bound forms of the TZF domain; and 2) NMR-based dynamic studies. Together, these studies should provide new insights into the mechanism of RNA recognition and, ultimately, mRNA deadenylation and decay mediated by the TTP-family of proteins.

Acknowledgments

This work was supported by the Intramural Research Program of the National Institute of Environmental Health Sciences, NIH. We are grateful to the Duke NMR Spectroscopy Center for use of instrumentation, data analysis, and training, and to Dr. Leonard Spicer for advice and encouragement. Thanks also to Geoffrey Mueller and Robin Stanley for helpful comments on the manuscript.

Footnotes

Conflict of interest

The authors declare that they have no conflict of interest.

Compliance with ethical standards

The authors declare that the experiments described herein comply with the current laws of the United States.

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