A diecast mineralization process forms the toughmantis shrimp dactyl clubShahrouz Aminia, Maryam Tadayona, Jun Jie Lokea, Akshita Kumara, Deepankumar Kanagavela, Hortense Le Ferranda,Martial Duchampb, Manfred Raidac, Radoslaw M. Sobotad, Liyan Chend, Shawn Hoone, and Ali Misereza,f,1

aCentre for Biomimetic Sensor Science, School of Materials Science and Engineering, Nanyang Technological University (NTU), 639798 Singapore; bSchool ofMaterials Science and Engineering, NTU, 639798 Singapore; cLife Science Institutes, Singapore Lipidomics Incubator, National University of Singapore (NUS),117456 Singapore; dFunctional Proteomics Laboratory, Institute for Molecular, Cell, and Development Biology, Agency for Science, Technology, andResearch (A*Star), 138673 Proteos, Singapore; eMolecular Engineering Laboratory, Biomedical Sciences Institutes, A*Star, 138673 Proteos, Singapore;and fSchool of Biological Sciences, NTU, 637551 Singapore

Edited by Lia Addadi, Weizmann Institute of Science, Rehovot, Israel, and approved March 19, 2019 (received for review October 2, 2018)

Biomineralization, the process by which mineralized tissues growand harden via biogenic mineral deposition, is a relatively lengthyprocess in many mineral-producing organisms, resulting in challengesto study the growth and biomineralization of complex hard miner-alized tissues. Arthropods are ideal model organisms to studybiomineralization because they regularly molt their exoskeletonsand grow new ones in a relatively fast timescale, providing oppor-tunities to track mineralization of entire tissues. Here, we monitoredthe biomineralization of the mantis shrimp dactyl club—a modelbioapatite-based mineralized structure with exceptional mechanicalproperties—immediately after ecdysis until the formation of the fullyfunctional club and unveil an unusual development mechanism. Aflexible membrane initially folded within the club cavity expands toform the new club’s envelope. Mineralization proceeds inwards bymineral deposition from this membrane, which contains proteins reg-ulating mineralization. Building a transcriptome of the club tissue andprobing it with proteomic data, we identified and sequenced ClubMineralization Protein 1 (CMP-1), an abundant mildly phosphorylatedprotein from the flexible membrane suggested to be involved in cal-cium phosphate mineralization of the club, as indicated by in vitrostudies using recombinant CMP-1. This work provides a comprehen-sive picture of the development of a complex hard tissue, from thesecretion of its organic macromolecular template to the formation ofthe fully functional club.

biomineralization | bioapatite | ecdysis | stomatopod dactyl club |mineralization proteins

Hard mineralized tissues grow through biogenic mineral de-position (biomineralization) and this process is a central

attribute of vertebrate development (1). However, investigatingthe growth process of entire hard tissues in vertebrates such asbone or teeth is challenging, owing to the relatively long timescaleover which mineralized tissues are formed (2, 3) and to sampleavailability. In contrast, crustaceans are convenient model organ-isms to study biomineralization because they regularly shed theirmineralized exoskeletons (cuticles) and grow new ones throughmolting cycles (4, 5). Specifically, molting and calcification of cu-ticles occur in just a few days or weeks, providing the distinctiveopportunity to follow the entire biomineralization process formodel organisms that can be maintained in the laboratory.Molting, the shedding (or ecdysis) of the exoskeleton, is an

essential event of arthropod development, during which the hardexoskeleton is replaced with a fresher, slightly larger one to ac-commodate the animal’s growth. Following molting, the freshlyformed exoskeleton is still soft and cannot fulfill its function,namely providing a protecting barrier against predators, pathogens,or the natural environment. Whereas molting takes just a few mi-nutes, mineralization of the new exoskeleton is longer, from days toweeks. Nevertheless, compared with vertebrate mineralization, theprocess is short enough such that the different stages can be studiedin the laboratory with convenient model organisms (5).

We used the dactyl club of stomatopods (mantis shrimps) as amodel structure to study the entire formation of hard and toughapatite-based mineralized appendages. The club is a biologicalhammer used by stomatopods to fracture the hard shells of theirpreys and has emerged in recent years as a fascinating modelstructure of bioinspired materials (6–10). The club is the mostmineralized appendage of the dactyl segment and exhibits acomplex architecture across multiple length scales, allowing theanimal to deliver extremely high impact forces against its targetswithout sustaining macroscopic fracture. In brief, the dactyl clubis a multilayer composite at the mesoscale that can be broadlyseparated into an outer region that expands toward the impactsurface and an inner bulk region. Both regions exhibit distinctchemical compositions and microstructures. The outer region ismostly made of crystalline fluorapatite (FAP) nanorods that arepreferentially oriented perpendicular to the impact surface, witha small presence of calcium sulfate (7). Moving toward the bulk,crystallinity of FAP decreases and the mineral phase graduallytransitions toward amorphous calcium phosphate (ACP). Theinner bulk region contains both ACP as well as amorphous cal-cium carbonate (ACC) that decorate chitin fibrils arranged in a

Significance

Monitoring hard tissues calcification using vertebrates is chal-lenging, owing to the internal location and slow biomineraliza-tion process of these tissues. Crustaceans are ideal modelorganisms to overcome this challenge because they regularlymolt their exoskeletons. Using the ultratough mantis shrimpdactyl club as a model biomineral, we detect all stages duringthe development of a calcified tissue, from secretion of theorganic template that regulates mineral deposition to matu-ration of the functional club. We unveil a peculiar growthmechanism: a flexible membrane initially folded in the clubcavity expands after ecdysis to form the new club outer en-velope from which biomineralization proceeds. A main phos-phorylated protein within that membrane is sequenced anddemonstrated to regulate mineral crystal growth.

Author contributions: S.A. and A.M. designed research; S.A., M.T., J.J.L., A.K., D.K., H.L.F.,M.R., R.M.S., L.C., and S.H. performed research; S.A., J.J.L., A.K., H.L.F., M.D., M.R., R.M.S.,L.C., S.H., and A.M. analyzed data; and S.A. and A.M. wrote the paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

Published under the PNAS license.

Data deposition: Transriptomic data of O. scyllarus dactyl club have been deposited inthe NCBI BioProject (accession no. PRJNA528158. Proteomic data have been deposited inthe jPOST Repository, https://repository.jpostdb.org (accession no. JPST000563), and in theProteomeXchange Consortium database (accession no. PXD013153).1To whom correspondence should be addressed. Email: ali.miserez@ntu.edu.sg.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1816835116/-/DCSupplemental.

Published online April 11, 2019.

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helicoidal pattern (6, 10). Collectively this multilayer design en-dows the club with exceptional tolerance against contact stresses(8) and serves as inspiration for the design of damage-tolerantbiocomposites (11, 12).

Molting Stages of the Mantis ShrimpMantis shrimps shed their exoskeletons a few times per year. Duringthis process, they are vulnerable to attacks from other predators(such as crabs, their congeners, or starfish) since their raptorialappendages are not functional for either hunting or defense pur-poses. To mitigate this drawback, in the premolt stage, mantisshrimps secure a nest by shattering rocks, shells, and corals and thencollect the broken pieces to build a protecting nesting cavity (SIAppendix, Fig. S1 A–C). During this process, the dactyl clubs areeroded on their impact surface due to high-energy hits against rock-solid targets (SI Appendix, Fig. S1 E and F), though they do nosustain catastrophic fracture. Subsequently during the molting pe-riod, stomatopods hide in this nest and avoid external contact.Our initial observations (SI Appendix, Movie S1) revealed that

during molting, the entire cuticle including the eye cups and thedactyl clubs were shed and buried in the sand for partial reuti-lization of nutrients (5). Fig. 1A is a picture of a mantis shrimpimmediately after molting next to the old cuticle. We found thatwhile the new exoskeletons were slightly larger than those beforemolting, the new dactyl clubs initially appeared as folded mem-branes that were much smaller than the shed dactyl clubs (exuviae,Fig. 1 B and C). However, these folded membranes expandedduring the first hour after ecdysis to form the outer shape of thenew dactyl clubs. Once expanded these membranes were largerthan the exuviae but soft and flexible (Fig. 1D) and were filledwith hemolymph fluid (“crustacean blood”).A closer look revealed that these folded membranes were

initially stored inside the internal cavity of the old clubs (Fig. 1 Band C) before their rapid expansion, which occurred within thefirst few minutes following ecdysis. These observations indicatedthat these membranes act as templates for club formation, andthat they must be folded and flexible to allow easy extrusionfrom the internal cavity of the old club. Energy dispersive X-rayspectroscopy (EDS) elemental mapping of a dactyl club cross-sectional cut (SI Appendix, Fig. S2A) confirmed the presence ofthe premolt membrane inside the internal cavity of the old dactyl

club, mostly composed of organic phases, as shown by the highcarbon content and the absence of calcium. We also note the pres-ence of sulfur in the premolt membrane, which may act as a reservoirfor calcium sulfate that is also found in the fully formed clubs (7).The fresh cuticle does not provide protection against external

threats: although the expanded membranes displayed the overallgeometry of a mature club, they were not functional due to theirweak mechanical properties (they could easily be bent and torn byhand), which explains why mantis shrimps refuse to hit any targetand hid inside their nest after ecdysis. Since their survival dependson a fully functional dactyl club, they must rapidly build a new one,thus providing a unique opportunity to study the entire bio-mineralization process. In our aquaria containing artificial seawater,we found that a partially functional club was formed within a week.

Formation of the Club by a Diecast MechanismWe followed our initial observations of the molting process withsystematic investigations of the different stages of dactyl clubformation right after ecdysis by examining the club structure andmechanical properties at different development stages, whichrevealed an unusual formation mechanism that, to the best ofour knowledge, has not been previously reported in highly min-eralized hard tissues. In clear contrast with the hypothesis of aninside-to-outward growth (13), we found that the club is built byan outside-to-inward mechanism as corroborated by optical andscanning electron microscopy (SEM) images of dactyl clubs atvarious development stages shown in Fig. 2. Within the first hourafter ecdysis, the folded membrane expanded to form the outergeometrical shape of the new club (Fig. 1D). This expandedmembrane was initially flexible but became a delicate andbrittle shell on the second day. We also observed the presenceof microchannels on day 2 (Fig. 2B, Middle Left and SI Ap-pendix, Fig. S2B) that likely enables the continuous growth ofthe outer layer by providing diffusion paths for the delivery ofionic species and/or nanocluster precursors (14) required formineral deposition.The formation of the inner, less mineralized layer of the club

started after a few days and included the deposition of nano-fibrils (Fig. 2 A and B, Middle Right) that form the helicoidalchitinous-based microstructure previously reported (6). After1 wk, the dactyl club became functional and the animals started

Fig. 1. Ecdysis process of the mantis shrimp. (A) Ecdysis of the entire mantis shrimp cuticle (Left), including the dactyl clubs (Right). The exuvia (shed cuticle) isseen next to the animal. (B) Exuvia and new appendages dissected from a mantis shrimp during ecdysis showing the folded membrane that was stored insidethe old dactyl club. (C) Micro-CT image of an old dactyl club illustrating the presence of the internal cavity inside the dactyl club. (D) Within an hour, thefolded membrane (Left) expands to take the shape of the new dactyl club (Middle), which is still soft and flexible (enlarged image Right) and contains thehemolymph fluid made mostly of hemocyanin (SI Appendix, Fig. S3 for proteomic analysis).

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venturing out of their nest to hunt for food and reengage instriking actions. Finally, after about 1 mo the dactyl clubs werefully formed and exhibited the complex multiscale architecturewith their characteristic high impact tolerance (6, 7) (Fig. 2 A andB, Right). These observations can be summarized in the schematiccartoon shown in Fig. 2D. The folded membrane rapidly expandsafter ecdysis, an unfolding process that determines the final outershape of the dactyl club. Club formation is then initiated from thisouter envelope and proceeds toward the inner layer, a mechanismresembling diecast processing. In the final stage, a new internalcavity of the club is maintained, which will serve as a template forthe formation of the next membrane and club.

Structural Formation and Mechanical StiffeningConcomitant to microscopic observations, we also probed thestructural and mechanical properties of the club at various de-velopment stages (from a few hours until 1 wk) using Ramanspectroscopy, EDS analysis, and nanomechanical measurements,and the data were compared with those from fully formed clubs.In the expanded membrane, only peaks associated with organicphases were detected at ∼1,160 cm−1 and 1,520 cm−1 (Fig. 3A),which can be assigned to proteins and chitin. On the other hand,no peaks related to calcium phosphate minerals were observed.However, 2 d after ecdysis the characteristic ν1 peak of apatite at965 cm−1 was detected with weak intensity, suggesting that min-eral deposition had been initiated. The ν1 peak intensity stronglyincreased after 1 wk and was the most prominent, with the overallspectrum similar to that of the fully formed clubs in the outer re-gion near the impact surface (Fig. 3A). We also noted the presenceof calcium sulfate (ν1 peak at 1,007 cm−1) previously found to be

colocalized with FAP crystallites (7), which was detected 2 d afterecdysis (Fig. 3A, Inset). EDS analysis confirmed the absence of Caand P in the premolt membrane and immediately after expansion(Fig. 3B), but both elements increased after 24 h with a Ca/P weightratio of ∼1.7, corresponding to calcium phosphate. In parallel weevaluated the mechanical properties of the club at various de-velopmental stages by nanoindentation and found that the ex-panded membrane in hydrated conditions was soft and flexible(elastic modulus E = 0.12 ± 0.03 GPa). After 48 h, the modulusstrongly increased by nearly two orders of magnitude (E = 10.2 ±2.2 GPa) and after 1 wk E reached 53.5 ± 3.6 GPa, which is ∼85%that of mature dactyl clubs (E = 62.8 ± 1.9 GPa).

In Vitro Apatite BiomineralizationSince biomineralization of the club was clearly initiated from theexpanded membrane, we hypothesized that the membrane con-tains the proteins that template and regulate apatite nucleationand growth during mineralization of the club. To test this hy-pothesis, we conducted an in vitro mineralization assay designedafter ref. 15 and incubated a membrane in a buffer saturatedwith Ca2+ and PO4

3− ions for 7 d (Fig. 4A), after which the in-cubated membrane was thoroughly rinsed and probed by SEM,EDS, Raman spectroscopy, and nanoindentation.We first verified the absence of calcium phosphate in the fresh

and rinsed expanded membrane by Raman spectroscopy (Fig.4C), which was further validated by EDS measurements thatconfirmed the absence of Ca (Fig. 4D). After incubation, Ramanspectra clearly indicated the presence of crystalline calcium phos-phate with the appearance of the intense PO4

3− ν1 vibrational peakat 965 cm−1. SEM observations showed the presence of spherical

Fig. 2. Structure of the dactyl club at various time points of the molting and formation process (premolt, and 2 d, 1–2 wk, and 1 mo after molting). (A)Optical micrographs of cross-sectional cuts of entire clubs. (B) Low-magnification (Upper) and higher-magnification (Lower) SEM micrographs. The club growsby an outside-to-inward mechanism the first month following ecdysis. It starts with expansion of the premolt membrane within the first hours followingecdysis to form the overall envelope of the new dactyl club. (C) Carbon elemental map of a mature club before molting, showing the presence of a premoltmembrane inside the internal club cavity. Adapted from ref. 7. (D) Schematic of the formation process of the dactyl club during the first month after ecdysis.

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particles at ∼100 nm on the fractured surface of the membrane(Fig. 4B). Point analysis of these particles by EDS (Fig. 4D)revealed a high intensity of Ca and P elements that corroboratedthe formation of calcium phosphate. Furthermore, the elasticmodulus (hydrated conditions) of the membrane increased up to20-fold after in vitro mineralization (Fig. 4E). As a control, theidentical incubation experiment was conducted on a membranethat was previously subjected to an alkali peroxidation treatmentto remove proteins, so that only chitin was left as the organicphase in the membrane (9). No calcium phosphate formation wasobserved after 7 d of incubation as evidenced by EDS measure-ments (SI Appendix, Fig. S4).In addition, we also found that the flexible membrane imme-

diately after molting was filled with the hemolymph fluid (Fig. 1D),which mostly comprises hemocyanin as confirmed by a compre-hensive set of proteomic analyses of the fluid (SI Appendix, Fig. S3).Previous studies have suggested that crustacean hemocyanin is notrestricted to its well-known oxygen-carrying role but may also beinvolved in calcium storage (16, 17) and transport to the growingcuticle (18). For example, we observed by transmission electronmicroscopy (TEM) dense amorphous granules ∼50–100 nm indiameter in the native hemolymph, which were enriched with Caand P (SI Appendix, Fig. S5). These granules may be condensedphosphates forming complexes with Ca2+ ions (19) that may serve

as P and Ca storage to accelerate club mineralization after molting.After incubating the hemolymph with Ca2+/PO4

3− saturated solu-tion and washing the samples, a high content of Ca and P remainedwithin these granules as evidenced by the strong signal intensity ofCa and P EDS peaks under identical acquisition conditions (SIAppendix, Fig. S6), possibly pointing out toward a storage role ofmineralization ions for the hemolymph. Additionally, the granuleswere amorphous, as no evidence of crystallinity was detected byelectron diffraction (SI Appendix, Fig. S6C).

Identification and Sequencing of Proteins ControllingApatite Nucleation and GrowthHaving established that proteins within the flexible membranecould regulate apatite nucleation and growth, we sought to iden-tify and sequence these putative mineralization proteins using acombined transcriptomic/proteomics strategy we previously de-veloped (20–22). First, transcriptome libraries (23) were assem-bled from mRNAs extracted from the epithelial cells of matureclubs of adult mantis shrimps as well as from larvae that werereared in our laboratory. Since there are no reference genomes forstomatopods, we generated transcript databases by de novo tran-script assembly with the Trinity software suite (24). In parallel, weextracted proteins from flexible membranes immediately afterecdysis using a guanidine thiocyanate-based buffer to maximizeprotein extraction. SDS polyacrylamide gel electrophoresis (SDS/PAGE) of the extracts revealed broad faint bands in the 15- to 20-kDa range as well as a sharp faint band between 60 and 75 kDa(Fig. 5A). These bands were excised from the gel and digestedwith trypsin. Subsequently, tryptic peptides were extracted fromthe gel slices and separated on a nano C18 high-performanceliquid chromatography (LC) column and subjected to LC tan-dem mass spectrometry. De novo sequencing of the tryptic pep-tides was obtained using the PEAKS studio 8.0 software (25). Denovo peptide fragments identified with PEAKS were then screeneddirectly against the transcriptome libraries of the club using theSpider and PEAKS search routines.We further narrowed down the transcriptome screening of

proteins detected by de novo sequencing with a combination ofthe following criteria: (i) molecular weight (MW) larger than50 kDa to match the main sharp band detected by SDS/PAGE;(ii) high transcript levels; (iii) enriched with acidic residues sincecalcium phosphate mineralization is well established to be con-trolled by highly acidic proteins (26, 27); and (iv) presence of chitin-binding domains, since the flexible membrane is a protein/chitincomplex. We identified five proteins containing chitin-bindingdomains in the club transcriptome (SI Appendix, Table S1 and afull list of tryptic peptides shown in SI Appendix, Table S2), one ofwhich was abundant with acidic residues [8.3 mol% aspartic acid(Asp) and 4.1 mol% of glutamic acid (Glu)], with no homology toany known protein, which we termed Club Mineralization Protein1 (CMP-1). We obtained the full-length sequence of CMP-1 byRACE-PCR (28) using the cDNA library of the club as a templatefor PCR, and the final gene was sequenced with Sanger se-quencing. Both the signal peptide as well as a stop codon in thegene encoding CMP-1 were detected, confirming that the full-length sequence of CMP-1 was achieved. The MW of CMP-1 is65 kDa, matching the MW of the sharp band detected by SDS/PAGE from the flexible membrane extract (Fig. 5A).To confirm that this band corresponded to CMP-1, we con-

ducted additional protein extraction from a flexible membraneusing a sonication method and ran the extract by SDS/PAGE (SIAppendix, Fig. S7A), followed by LC MS/MS analysis of individualexcised bands (29). CMP-1 tryptic peptides were identified (SIAppendix, Table S3) with a protein coverage of 39% (SI Appendix,Fig. S7C), therefore corroborating the assignment of CMP-1 tothe sharp band between 50 and 75 kDa. All detected peptideswere from the N terminus due to the absence of cleavage sites inthe C terminus. Furthermore, we found by using quercetin-based

Fig. 3. Structural and mechanical properties of the dactyl club at differentdevelopment stages. (A) Raman spectra at different stages of formation. FAP(ν1 vibration mode at 965 cm−1) is not detected in the expanded membrane,weakly observed in the 24- to 48-h postmolt club, and abundant in the 7-dpostmolt club. (B) EDS point analysis at different club formation stages(variation in Ca from 24-h postmolt onward is attributed to statistical vari-ability between different animals). (C) Elastic modulus (E) of the club (outerregion) at different stages of formation. The expanded membrane in thehydrated state is soft (E = 0.12 ± 0.03 GPa). Two days after ecdysis (newlyformed club), E increases to ∼10.2 ± 2.2 GPa. After 2 wk (immature club) E =53.5 ± 3.6 GPa, close to E in mature clubs (E = 62.8 ± 1.9 GPa).

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phosphostain (SI Appendix, Fig. S7B) that CMP-1 was mildlyphosphorylated, and two phosphorylated Ser residues (S55 andS226) were detected by LC MS/MS with high confidence (SIAppendix, Fig. S7 C and D). CMP-1 harbors several sequencecharacteristics that make it particularly suitable as an organictemplate for biocomposite mineralization. First, it features a centraldomain (R294–Y339) with sequence homology to the chitin-binding4 superfamily (Fig. 5B), which was predicted to bind to chitin, basedon molecular dynamic (MD) simulations (Materials and Methodsand SI Appendix, Fig. S8). Second, it contains a very acidic N-terminal domain that we hypothesized could bind Ca2+ ions tocontrol calcium phosphate nucleation. This dual binding role to bothchitin nanofibrils (to form the organic template of the flexible mem-brane) and to Ca2+ for subsequent mineral deposition is reminiscentof GAP65 identified in the gastrolith of the crayfish that stabilizesamorphous calcium carbonate (30). Notably, the N-terminal do-main contains two poly-Asp rich motifs that are strikingly similar topoly-Asp domains found in osteopontin, a main protein regulatingapatite mineralization in bone (31, 32). The first motif (D154–D175)is 22 amino acid long and contains 17 Asp residues with the longeststretch comprising 10 Asp. The second motif (D229–D246) is

17 amino acid long and is comprised of 14 Asp intervened withthree Phe residues. Similar to osteopontin and other proteins in-volved in biomineralization (31), it is noteworthy that CMP-1 isalso phosphorylated, although to a much lesser extent with onlytwo detected sites at S55 and S226. Additional phosphorylatedsites may also be present in the C terminus; however the lack ofcleavable tryptic peptides in this region precluded us fromidentifying them.

In Vitro Apatite Mineralization and Microdroplet Formationfrom Recombinant CMP-1To confirm our hypothesis that CMP-1 can nucleate and regulateapatite formation, we cloned, expressed, and purified CMP-1 tohomogeneity (Fig. 5C) and then conducted in vitro assays byincubating soluble recombinant CMP-1 (rCMP-1) in a Ca2+/PO4

3− saturated solution using similar protocols as for the nativemembrane. After 6 d of incubation, we observed rod-like min-erals by SEM (Fig. 5D), which were composed mainly of Ca andP as inferred from EDS, with a Ca/P ratio of 1.72, close to thevalue (1.68) documented for crystalline apatite. To gain furtherdetails on the nucleation and growth of these rods, we conducted

Fig. 4. In vitro mineralization studies using the native proteins of the dactyl club as template for biomineralization. (A) Expanded membrane forming theouter envelope of the club by incubation of the flexible extended membrane in buffer saturated with Ca2+ and PO4

3− ions. (B) Pre- and postincubation SEMimages revealing the flexible and fibrous structure of the expanded membrane, which turned into a rigid structure after incubation in Ca2+/PO4

3− saturatedsolution. (C and D) Raman (C) and EDS (D) spectra of an expanded membrane and a membrane incubated in Ca2+/PO4

3− saturated solution. No trace ofmineralization in the extended membrane was detected (no PO4

3− band in the Raman spectrum, and no Ca peak in the EDS spectrum). The incubatedmembrane, on the other hand, was mineralized as indicated by the presence of PO4

3− bands (ν1, ν2, and ν4 in the Raman spectrum) and by the strong intensityof Ca and P peaks in the EDS spectrum. (E) The elastic modulus (E) of the membrane (hydrated conditions) increased up to 20-fold after 1 wk of incubation inthe Ca2+/PO4

3− saturated solution from E = 0.12 ± 0.03 GPa to E = 2.16 ± 0.3 GPa, and from E = 0.16 ± 0.03 GPa to E = 1.44 ± 0.3 GPa, for samples 1 and 2,respectively.

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TEM measurements at various time points (Fig. 5E). At 0.5 dand 1 d, we observed nanoclusters 40–60 nm in size, and theseclusters were amorphous, based on their electron diffractionpatterns (Fig. 5E, Bottom). After 1.5 d, larger and elongated rodswere observed, which were oriented crystalline apatite as evi-denced by electron diffraction. After 6 d of incubation, thecrystallinity of the apatite rods further increased as revealed bysingle diffraction spots on the electron diffraction pattern (forfull assignment of diffraction peaks, see SI Appendix, Fig. S9).These data corroborate previous studies by Dey et al. (14) inwhich the amorphous-to-crystalline transition of calcium phos-phate was shown to be initiated through prenucleation ofnanoclusters. Among the invertebrates, we are only aware of oneother sequenced protein (from the crayfish molar tooth) thatalso regulates apatite formation (33). We emphasize that thesein vitro experiments were conducted using rCMP-1, which lacksphosphorylation detected in the wild-type CMP-1. Phosphory-lated proteins in biomineralization have been shown to stabilizeACP and inhibit crystallization in some cases (34, 35) or promoteapatite formation in others (36). Since ACP nanoclusters were

detected in our experiments before they crystallized into ori-ented apatite nanorods, this suggests that CMP-1 can both nu-cleate transient ACP and then promote apatite crystallizationregardless of phosphorylation. Furthermore, crystalline apatitewas detected early during club formation (Fig. 3A) in the outerlayer of the club (Fig. 2D), whereas ACP is not found in this layerof the club (7). Similarly in our in vitro mineralization experi-ments using the extracted flexible membrane as a template,crystalline apatite was formed after a few days of incubation butnot ACP (Fig. 4C). Since CMP-1 is abundant in the flexiblemembrane, the colocalization of crystalline apatite and CMP-1 suggests that the role of phosphorylation in CMP-1 may beto regulate the kinetics of apatite crystallization, but this hy-pothesis remains to be validated. To answer this question, it willbe critical to determine the currently unknown in vivo conditions(such as pH and ionic strength) under which the club is formedbecause they likely influence protein activity (37).To assess the expression level of CMP-1, we also conducted

qPCR experiments using mRNA extracted at different stages ofmolting. We found that the expression level increased from a few

Fig. 5. CMP-1 sequence and in vitro mineralization. (A) SDS/PAGE of protein extract from the flexible membrane. The arrow indicates CMP-1 as confirmed byMS/MS analysis of individual bands conducted on additional gel extracts (SI Appendix, Fig. S7A). This band also stained positively for the quercetin phos-phostain (SI Appendix, Fig. S7B) indicating that CMP-1 is mildly phosphorylated. (B) Full-length sequence of CMP-1, obtained by searching flexible membranepeptides identified by MS/MS against the club tissue transcriptome, followed by RACE-PCR. *Phosphorylation sites detected by MS/MS. Bioinformatic predictionsare underlined in yellow (β-sheets) and blue (α-helices), or highlighted in green (chitin-binding domain). Asp-rich cluster regions that likely bind Ca2+ ions arehighlighted in red. The structure of the chitin-binding domain computed byMD simulations is also shown below (see SI Appendix, Fig. S8 for more details). (C) SDS/PAGE of rCMP-1 after purification. (D) SEM image of rCMP-1 after 6 d of incubation in the Ca2+ and PO4

3− saturated solution (Top) and EDS spectrum (Bottom)showing the presence of rod-like structures containing P and Ca (Na, Mg, Al, and Si are from the soda-lime glass substrate). (E) TEM imaging (Top) and electrondiffraction patterns (Bottom) of rCMP-1 incubated in the Ca2+ and PO4

3− saturated solution at various time points. Amorphous nanoclusters containing Ca and Pwere observed at 0.5 d and 1 d. From 1.25 d and beyond crystalline apatite was observed, with increasing crystallinity over time. Full assignment of diffractionspeaks is shown in SI Appendix, Fig. S9.

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hours after molting to reach its highest level at day 2, and thendropped at day 7 (SI Appendix, Fig. S10). Since the outer (im-pact) layer of the club made of crystalline apatite is fully formedwithin the first few days after molting, and since the remainingformation of the club mostly involves deposition of amorphouscalcium carbonate within the interior of the club cavity (but noadditional crystalline apatite from day 7 onwards, see Fig. 2 Aand D), these results further support the notion that CMP-1plays an important role in regulating apatite crystallization.This is also in agreement with the fact that transcript levels(normalized for sequencing depth) were approximately one or-der of magnitude higher in the mature club compared with thesaddle segment of the dactyl appendage (SI Appendix, Table S1),which is made of amorphous calcium phosphate but does notcontain crystalline apatite (9), suggesting a direct correlationbetween calcium phosphate crystallinity and CMP-1 expressionlevel. Furthermore, transcript levels in whole larvae of mantisshrimps—which do not develop a club yet—were even lower,namely 500-fold less than in the club tissue.In addition to regulating apatite nucleation and growth, we

also observed that rCMP-1 formed liquid droplets at pH 8.2 inthe presence of 5 mM CaCl2 and above, with droplet size in therange of 1–3 μm as revealed by SEM imaging and dynamic lightscattering measurements (SI Appendix, Fig. S11). This observationparallels recent findings by Bahn et al. (38) for Pfam-80 proteinthat controls calcium carbonate formation in oyster nacre, andfurther supports the growing concept that biomacromolecules con-trolling biomineralization phase separate into coacervate dropletsthat can sequestrate and concentrate inorganic ions for subsequentmineral deposition (38, 39). There has been mounting evidence inrecent years that proteins undergoing simple component liquid–liquid phase separation (coacervation) are intrinsically disordered(40) or contain intrinsically disordered regions (IDRs) (41).Inspecting CMP-1’s primary structure, we note that the C-terminalregion is highly enriched in glycine (Gly) and alanine (Ala) aminoacids and also contains a significant amount of proline (Pro), whichare characteristic features of IDRs as also confirmed by our bio-informatic predictions of this region (Fig. 5B). This suggests a rolefor the C terminus of CMP-1 to induce coacervation as a way toconcentrate inorganic ions before mineralization.It is very likely that other proteins detected in our combined

transcriptomic/proteomic dataset are also involved in club miner-alization since it is now well established that tissue biomineraliza-tion and crystal polymorphism is carefully orchestrated by multiplebiomacromolecules, including low MW compounds (42). Forexample, we found chitin-binding proteins with homology tocuticular proteins from other crustaceans, such as the Americanlobster or the brown crab (SI Appendix, Table S1), suggestingthat the flexible membrane is a macromolecular complex com-prising chitin and multiple chitin-binding proteins. Likewise,there are several other peptides uncovered by de novo MS/MSpeptide sequencing that are enriched with acidic residues (SIAppendix, Table S2) and that may also play a role in binding Caions and regulating apatite mineralization. Finally, phosphorylatedproteins with lower MWs than CMP-1 were also detected in theflexible membrane (SI Appendix, Fig. S7B), which is anothercharacteristic feature of proteins controlling biomineral formation(31). However, CMP-1 was the only protein we detected thatharbored several clusters of acidic residues (poly-Asp) along itsprimary sequence, which is a common feature associated withproteins controlling apatite formation. A comprehensive proteo-mic analysis of all proteins detected in the flexible membrane iscurrently underway and will be reported elsewhere.

ConclusionExploiting the fast molting process of crustaceans’ exoskeletons,we have unveiled the entire formation of the mantis shrimpdactyl club from ecdysis to the mature, impact-resistant club. Our

study reveals a peculiar development mechanism of the club. Theinner cavity of the fully formed club contains a folded flexiblemembrane that rapidly expands immediately after ecdysis toform the outer envelope of the club. Club mineralization ensuesvia an outside-to-inward mineral deposition mechanism, formingthe functional, impact-resistant club within a few weeks. The flexibleouter membrane is a chitin/protein macromolecular complex whoseproteins can trigger calcium phosphate nucleation and mineral-ization. Proteins within the flexible membrane were extractedfrom the membrane and sequenced via a dual transcriptomic/proteomic approach. One of the most abundant proteins (CMP-1)was in particular identified and its full-length amino acid sequenceobtained. CMP-1 comprises two key domains conferring the criticaldual function of binding to chitin to form the organic matrix, as wellas to Ca2+ ions to regulate calcium phosphate mineralization. Inaddition, CMP-1 is slightly phosphorylated and can phase separateinto liquid droplets, which are characteristics that may also play arole in the growth kinetics and polymorphism control of calciumphosphate during club formation. In vitro assays in the presenceof rCMP-1 resulted in the formation of apatite nanorods, sug-gesting the ability of CMP-1 to regulate crystalline apatite. Re-vealing the natural fabrication process by which a remarkabletough biomineralized appendage is produced offers bioinspiredlessons that may be applied in additive manufacturing of bioceramics,with potential applications for the next generation of orthopedicor dental implants.

Materials and MethodsDetailed experimental procedures are provided in SI Appendix,Materials andMethods. In brief, mantis shrimps (Odontodactylus scyllarus) were purchasedfrom aquarium suppliers in Singapore, maintained in artificial seawateraquaria, and carefully monitored to detect molting events. Dactyl clubs werecollected at various time points during molting and formation cycles, fromimmediately (flexible membrane) up to 1 mo (mature club) after moltingand used for subsequent studies. Biomineral characterization was carriedout by Raman confocal microspectroscopy, EDS, and optical microscopy.Mechanical characterization was carried out on hydrated samples usingdepth-sensing nanoindentation. For in vitro biomineralization using nativeflexible membranes, the latter were incubated in a Ca2+ and PO4

3− saturatedbuffer and analyzed postmineralization by Raman spectroscopy, EDS, andnanoindentation. Transcriptome libraries of dactyl appendage segments[namely the club (6–8) and the saddle (9)] as well as of whole larvae reared inour artificial seawater aquaria were prepared from mRNA extracted fromtissues that were initially stored in RNAlater solution. The libraries were se-quenced on a HiSEq. 2000 Illumina sequencer and the de novo transcript as-sembly was performedwith Trinity (24). For CMP-1 identification and sequencing,proteins were extracted from the flexible membrane and analyzed by SDS/PAGE, followed by LC MS/MS of both entire gels and excised individualbands. In both cases, trypsin in gel digestion was carried out, tryptic peptideswere analyzed with either PEAKS (25) or Proteome Discover 2.2 (Thermo),and then searched against our de novo transcriptome libraries. Full-lengthprotein sequencing of CMP-1 was achieved using both 3′ and 5′ RACE-PCRwith RNA extracted from the club tissue as the template. Gene encodingCMP-1 was cloned into pET28A plasmid, transformed to Escherichia coliBL21, and rCMP-1 was expressed in LB medium using isopropyl β-D-1-thio-galactopyranoside to induce expression. rCMP-1 was purified by immobilizedmetal affinity chromatography followed by FPLC and desalted before usage.In vitro mineralization assays were conducted by incubating rCMP-1 with aCa2+ and PO4

3− saturated buffer and analyzed by SEM, EDS, and TEM in boththe imaging and electron diffraction modes. Liquid–liquid phase separation(microdroplet formation) of rCMP-1 was achieved by pipetting the purifiedprotein in a CaCl2-containing Tris buffer. Secondary structure predictions andsequence alignment of CMP-1 were conducted using a range of bioinformatictools, including PSIPRED (43), BLASTp (44), CLUSTAL W (45), and Modeler (46).MD simulations of CMP-1 were carried out using Amber 12 (47). Chitin-CMP-1 interactions were modeled using a 3D structure of chitin from ChemSpiderand Schrodinger 9.0 with previously used protocols (48).

ACKNOWLEDGMENTS. We thank Isaiah Chua for helping with flexiblemembrane sample collection; Kong Kiat Whye for technical support withtranscriptome library preparation and qPCR measurements; and the centralFacilities for Analysis, Characterization, Testing, and Simulation at NTU for

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access to their electron microscopy facilities. This work was funded bythe Singapore National Research Foundation (NRF) through an individualNRF Fellowship (to A.M.), and by the Strategic Initiative on Biomimetic and

Sustainable Materials (IBSM, NTU). H.L.F. gratefully acknowledges the SwissNational Science Foundation for an individual postdoctoral scholarship(Grant_P2EZP2_172169). R.M.S. and L.C. were supported by A*Star core funding.

1. Weiner S, Addadi L (2011) Crystallization pathways in biomineralization. Annu RevMater Res 41:21–40.

2. Fincham AG, et al. (2000) Enamel biomineralization: The assembly and disassembly ofthe protein extracellular organic matrix. Development, Function and Evolution ofTeeth, eds Teaford MF, Ferguson MWJ, Meredith Smith M (Cambridge Univ Press,Cambridge, UK), pp 37–62.

3. Fratzl P, Fratzl-Zelman N, Klaushofer K, Vogl G, Koller K (1991) Nucleation andgrowth of mineral crystals in bone studied by small-angle X-ray scattering. CalcifTissue Int 48:407–413.

4. Bentov S, Abehsera S, Sagi A (2016) The mineralized exoskeletons of crustaceans.Extracellular Composite Matrices in Arthropods, eds Cohen E, Moussian B (SpringerInternational Publishing, New York), pp 137–163.

5. Luquet G (2012) Biomineralizations: Insights and prospects from crustaceans. ZooKeys176:103–121.

6. Weaver JC, et al. (2012) The stomatopod dactyl club: A formidable damage-tolerantbiological hammer. Science 336:1275–1280.

7. Amini S, et al. (2014) Textured fluorapatite bonded to calcium sulphate strengthenstomatopod raptorial appendages. Nat Commun 5:3187.

8. Amini S, Tadayon M, Idapalapati S, Miserez A (2015) The role of quasi-plasticity in theextreme contact damage tolerance of the stomatopod dactyl club. Nat Mater 14:943–950.

9. Tadayon M, Amini S, Masic A, Miserez A (2015) The mantis shrimp saddle: A biologicalspring combining stiffness and flexibility. Adv Funct Mater 25:6437–6447.

10. Yaraghi NA, et al. (2016) A sinusoidally architected helicoidal biocomposite. AdvMater 28:6835–6844.

11. Studart AR (2016) Additive manufacturing of biologically-inspired materials. ChemSoc Rev 45:359–376.

12. Amini S, Miserez A (2013) Wear and abrasion resistance selection maps of biologicalmaterials. Acta Biomater 9:7895–7907.

13. Guarín-Zapata N, Gomez J, Yaraghi N, Kisailus D, Zavattieri PD (2015) Shear wavefiltering in naturally-occurring Bouligand structures. Acta Biomater 23:11–20.

14. Dey A, et al. (2010) The role of prenucleation clusters in surface-induced calciumphosphate crystallization. Nat Mater 9:1010–1014.

15. Takeuchi A, et al. (2003) Deposition of bone-like apatite on silk fiber in a solution thatmimics extracellular fluid. J Biomed Mater Res A 65:283–289.

16. Adachi K, Endo H, Watanabe T, Nishioka T, Hirata T (2005) Hemocyanin in the exo-skeleton of crustaceans: Enzymatic properties and immunolocalization. Pigment CellRes 18:136–143.

17. Glazer L, et al. (2013) Hemocyanin with phenoloxidase activity in the chitin matrix ofthe crayfish gastrolith. J Exp Biol 216:1898–1904.

18. Ziegler A, Fabritius H, Hagedorn M (2005) Microscopical and functional aspects ofcalcium-transport and deposition in terrestrial isopods. Micron 36:137–153.

19. Omelon S, Habraken W (2016) Influence of condensed phosphates on the physicalchemistry of calcium phosphate solids. Inorganic Polyphosphates in Eukaryotic Cells,eds Kulakovskaya T, Pavlov E, Dedkova EN (Springer International Publishing, NewYork), pp 177–205.

20. Guerette PA, et al. (2013) Accelerating the design of biomimetic materials by in-tegrating RNA-seq with proteomics and materials science. Nat Biotechnol 31:908–915.

21. Loke JJ, Kumar A, Hoon S, Verma C, Miserez A (2017) Hierarchical assembly of toughbioelastomeric egg capsules is mediated by a bundling protein. Biomacromolecules18:931–942.

22. Tan Y, et al. (2015) Infiltration of chitin by protein coacervates defines the squid beakmechanical gradient. Nat Chem Biol 11:488–495.

23. Amini S, et al. (2019) Odontodactylus scyllarus transcriptome and gene expression.BioProject. Available at https://www.ncbi.nlm.nih.gov/bioproject?LinkName=biosample_bioproject&from_uid=11175071. Deposited March 20, 2019.

24. Grabherr MG, et al. (2011) Full-length transcriptome assembly from RNA-Seq datawithout a reference genome. Nat Biotechnol 29:644–652.

25. Ma B, et al. (2003) PEAKS: Powerful software for peptide de novo sequencing bytandem mass spectrometry. Rapid Commun Mass Spectrom 17:2337–2342.

26. Veis A (2005) Materials science. A window on biomineralization. Science 307:1419–1420.

27. Suzuki M, et al. (2009) An acidic matrix protein, Pif, is a key macromolecule for nacreformation. Science 325:1388–1390.

28. Scotto-Lavino E, Du G, Frohman MA (2006) 5′ end cDNA amplification using classicRACE. Nat Protoc 1:2555–2562.

29. Amini S, et al. (2019) Odontodactylus scyllarus club mineralization proteomics data(LC MS/MS). jPOST Repository. Available at https://repository.jpostdb.org/entry/JPST000563).Deposited March 6, 2019.

30. Shechter A, et al. (2008) A gastrolith protein serving a dual role in the formation of anamorphous mineral containing extracellular matrix. Proc Natl Acad Sci USA 105:7129–7134.

31. George A, Veis A (2008) Phosphorylated proteins and control over apatite nucleation,crystal growth, and inhibition. Chem Rev 108:4670–4693.

32. Sodek J, Ganss B, McKee MD (2000) Osteopontin. Crit Rev Oral Biol Med 11:279–303.33. Tynyakov J, et al. (2015) A crayfish molar tooth protein with putative mineralized

exoskeletal chitinous matrix properties. J Exp Biol 218:3487–3498.34. Glazer L, et al. (2010) A protein involved in the assembly of an extracellular calcium

storage matrix. J Biol Chem 285:12831–12839.35. Holt C, Wahlgren NM, Drakenberg T (1996) Ability of a beta-casein phosphopeptide

to modulate the precipitation of calcium phosphate by forming amorphous dicalciumphosphate nanoclusters. Biochem J 314:1035–1039.

36. He G, et al. (2005) Phosphorylation of phosphophoryn is crucial for its function as amediator of biomineralization. J Biol Chem 280:33109–33114.

37. Tsutsui N, Ishii K, Takagi Y, Watanabe T, Nagasawa H (1999) Cloning and expressionof a cDNA encoding an insoluble matrix protein in the gastroliths of a crayfish. ZoolSci 16:619–629.

38. Bahn SY, Jo BH, Choi YS, Cha HJ (2017) Control of nacre biomineralization by Pif80 inpearl oyster. Sci Adv 3:e1700765.

39. Ibsen CJS, Gebauer D, Birkedal H (2016) Osteopontin stabilizes metastable states priorto nucleation during apatite formation. Chem Mater 28:8550–8555.

40. Banani SF, Lee HO, Hyman AA, Rosen MK (2017) Biomolecular condensates: Orga-nizers of cellular biochemistry. Nat Rev Mol Cell Biol 18:285–298.

41. Brangwynne Clifford P, Tompa P, Pappu Rohit V (2015) Polymer physics of intracel-lular phase transitions. Nat Phys 11:899–904.

42. Sato A, et al. (2011) Glycolytic intermediates induce amorphous calcium carbonateformation in crustaceans. Nat Chem Biol 7:197–199.

43. McGuffin LJ, Bryson K, Jones DT (2000) The PSIPRED protein structure predictionserver. Bioinformatics 16:404–405.

44. Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ (1990) Basic local alignmentsearch tool. J Mol Biol 215:403–410.

45. Thompson JD, Higgins DG, Gibson TJ (1994) CLUSTAL W: Improving the sensitivity ofprogressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res 22:4673–4680.

46. �Sali A, Blundell TL (1993) Comparative protein modelling by satisfaction of spatialrestraints. J Mol Biol 234:779–815.

47. Pearlman DA, et al. (1995) AMBER, a package of computer programs for applyingmolecular mechanics, normal mode analysis, molecular dynamics and free energycalculations to simulate the structural and energetic properties of molecules. ComputPhys Commun 91:1–41.

48. Kannan S, et al. (2015) Probing the binding mechanism of Mnk inhibitors by dockingand molecular dynamics simulations. Biochemistry 54:32–46.

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