, 20130339, published 15 May 2013280 2013 Proc. R. Soc. B Belikov, Dorte Janussen, Vasilii V. Bazhenov and Gert WörheideN. Sivkov, Denis Vyalikh, René Born, Thomas Behm, Andre Ehrlich, Lubov I. Chernogor, SergeiTabachnick, Micha Ilan, Allison Stelling, Roberta Galli, Olga V. Petrova, Serguei V. Nekipelov, Victor Hermann Ehrlich, Oksana V. Kaluzhnaya, Mikhail V. Tsurkan, Alexander Ereskovsky, Konstantin R. First report on chitinous holdfast in sponges (Porifera)  

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ResearchCite this article: Ehrlich H, Kaluzhnaya OV,

Tsurkan MV, Ereskovsky A, Tabachnick KR, Ilan

M, Stelling A, Galli R, Petrova OV, Nekipelov SV,

Sivkov VN, Vyalikh D, Born R, Behm T, Ehrlich

A, Chernogor LI, Belikov S, Janussen D,

Bazhenov VV, Worheide G. 2013 First report

on chitinous holdfast in sponges (Porifera).

Proc R Soc B 280: 20130339.

http://dx.doi.org/10.1098/rspb.2013.0339

Received: 11 February 2013

Accepted: 18 April 2013

Subject Areas:biochemistry, evolution, structural biology

Keywords:chitin, Porifera, holdfast, endemic sponges

Authors for correspondence:Hermann Ehrlich

e-mail: hermann.ehrlich@physik.tu-freiberg.de

Gert Worheide

e-mail: woerheide@lmu.de

Electronic supplementary material is available

at http://dx.doi.org/10.1098/rspb.2013.0339 or

via http://rspb.royalsocietypublishing.org.

& 2013 The Author(s) Published by the Royal Society. All rights reserved.

First report on chitinous holdfastin sponges (Porifera)

Hermann Ehrlich1, Oksana V. Kaluzhnaya3, Mikhail V. Tsurkan4,Alexander Ereskovsky5, Konstantin R. Tabachnick6, Micha Ilan7,Allison Stelling8, Roberta Galli8, Olga V. Petrova9, Serguei V. Nekipelov9,Victor N. Sivkov9, Denis Vyalikh10, Rene Born11, Thomas Behm1,Andre Ehrlich2, Lubov I. Chernogor3, Sergei Belikov3, Dorte Janussen12,Vasilii V. Bazhenov1 and Gert Worheide13,14,15

1Institute of Experimental Physics, and 2Institute of Mineralogy, TU Bergakademie Freiberg,09599 Freiberg, Germany3Limnological Institute SB RAS, 664033 Irkutsk, Russia4Max Bergmann Centre for Biomaterials, Leibniz Institute of Polymer Research, 01062 Dresden, Germany5Mediterranean Institute of Biodiversity and Ecology, CNRS, IRD, Aix-Marseille University,13007 Marseille, France6P.P. Shirshov Institute of Oceanology, Russian Academy of Sciences, Moscow, Russia7Department of Zoology, Tel Aviv University, Tel Aviv 69978, Israel8Carl-Gustav-Carus Klinikum, TU Dresden, 01307 Dresden, Germany9Department of Mathematics, Komi SC UrD RAS, Syktyvkar, Russia10Institute of Solid State Physics, TU Dresden, 01062 Dresden, Germany11Institute of Materials Science, TU Dresden, 01062 Dresden, Germany12Forschungsinstitut und Naturmuseum Senckenberg, 60325 Frankfurt am Main, Germany13Department of Earth and Environmental Sciences, Palaeontology & Geobiology, and 14GeoBio-Center,Ludwig-Maximilians-Universitat Munchen, Richard-Wagner-Strasse 10, 80333 Munchen, Germany15Bayerische Staatssammlung fur Palaontologie und Geologie, Richard-Wagner-Strasse 10,80333 Munchen, Germany

A holdfast is a root- or basal plate-like structure of principal importance that

anchors aquatic sessile organisms, including sponges, to hard substrates. There

is to date little information about the nature and origin of sponges’ holdfasts in

both marine and freshwater environments. This work, to our knowledge,

demonstrates for the first time that chitin is an important structural component

within holdfasts of the endemic freshwater demosponge Lubomirskia baicalensis.Using a variety of techniques (near-edge X-ray absorption fine structure,

Raman, electrospray ionization mas spectrometry, Morgan–Elson assay and Cal-

cofluor White staining), we show that chitin from the sponge holdfast is much

closer to a-chitin than to b-chitin. Most of the three-dimensional fibrous skeleton

of this sponge consists of spicule-containing proteinaceous spongin. Intriguingly,

the chitinous holdfast is not spongin-based, and is ontogenetically the oldest part

of the sponge body. Sequencing revealed the presence of four previously unde-

scribed genes encoding chitin synthases in the L. baicalensis sponge. This

discovery of chitin within freshwater sponge holdfasts highlights the novel and

specific functions of this biopolymer within these ancient sessile invertebrates.

1. IntroductionEnvironmental characteristics are known to influence the gross morphology of many

benthic organisms including sponges (Porifera; [1,2]). This variability in shape is the

consequence of both strategic (during a long-term evolutionary response to environ-

mental pressures) and tactical (in response to local environmental pressures)

processes. At local scales, morphology correlates with wave action, current flow

rate and sedimentation for both a number of sponge species and entire sponge

assemblages [3–6]. Wave action and current flow rate have direct influences not

only on sponge morphology, but also on the strength of attachment to substrata

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by its holdfast. For example, wave action is known to affect

sponge morphological types, as many of the more delicately

branching species are destroyed by drag [4,5]. Therefore, the

limits of morphological adaptation for any particular sponge

species may reduce species richness at sites of high wave exposure

flow. Wave action removes delicate sponge forms such as the ped-

unculate and arborescent shapes. Encrusting and robust species

are more suited to this environment [7]. However, the biomecha-

nical basis for such morphological changes has rarely been

documented [5]. The holdfasts of sponges that live in muddy

substrates often have complex tangles of root-like growths, how-

ever the holdfasts of organisms that live on smooth surfaces

(such as the surface of a boulder) have the base of the holdfast

literally glued to the surface. For example, the globular demos-

ponge Cinachyra subterranea, van Soest & Sass [8] has been

found on vertical walls and the floors of caves flooded with

water at marine salinity levels [8]. Attachment to the rock is

accomplished not by a root of spicules, but by a smooth, flat,

disk-like holdfast. Some sponges, e.g. species of Tentorium,

may show adaptation to both soft bottom and hard substrate [9].

Detailed analysis of the literature with regard to the

nature and origin of the poriferan holdfast suggests that it

is a complex structure that is initially developed by the

larvae. When a sponge begins its life cycle, it is a microscale

free-swimming larva in the water column. The tiny sponge

must settle down on a substrate and establish a niche for

itself to survive. After settlement, the larvae metamorphoses

and begin to transform their organization into the adult body

plan. During this process, the outermost layer of cells (the

pinacoderm) covers the metamorphosing sponge, as well as

the adult body [10]. The basal pinacoderm (basopinacocytes)

secretes a mixture of spongin (collagen-like protein) and com-

plex carbohydrates (probably in the form of a fibrillar

spongin–polysaccharide complex) that allows the animal to

attach to a substrate [11]. The spongin attachment plaques

can be seen as the precursor of the sponge holdfast: it is

secreted by basopinacocytes, and the protein–carbohydrate-

based glue secreted by these cells holds the sponge in place.

According to the traditional point of view, spongin is the

basic component of the sponges’ organic skeleton. Taxono-

mically, spongin is a character of the class Demospongiae,

which comprises the highest number of known species (cur-

rently more than 8000). Only the representatives of marine

demosponge Order Verongida possess skeletons that consist

mostly (up to 70%) of chitin, and not spongin [12,13].

It was proposed [14] that spongin sticks the animal to its sub-

stratum [15], links its skeletal spicules together, and is also

present within the coat of sponge gemmules [16]. Although the

spongin matrix has been defined as an exoskeleton [16], spongins

exhibit different morphological aspects among demosponges

and vary according to the tissues. It is currently not known if

all spongin assemblies are equivalent [11,17], or whether or

not they are entirely made of short-chain sponge collagens [14].

Thus, according to traditional point of view the reticulate

skeleton in most demosponges arises from the basal spongin

plate [18] and their spicules (if present) are cemented by varied

amounts of spongin in the form of bundles and networks.

Formation of the spongin attachment plaques (¼ anchoring

layer, basal layer and spongin lamella) as the possible precursor

of the sponge holdfast has been investigated during aggregate

differentiation in demosponge cells [15] and for settlement

and metamorphosis of the parenchymella larvae [19,20] for

freshwater sponges, including the metamorphed larva stage.

According to observations by transmission electron

microscopy and scanning electron microscopy, fibrous

material is found within the basal plates of young sponges,

and within the holdfasts of adult ones. Intriguingly, to our

best knowledge, to date no reports exist containing detailed

bioanalytical investigations confirming that the fibrous

material observed is really collagen-like spongin. However,

from a methodological point of view, verification of the pres-

ence of spongin within skeletal formations is very simple.

This arises from the excellent solubility of spongin in alkaline

solutions. This property of spongin is well known, and was

first described by Kunike [21]. Our preliminary investigations

attempted to isolate peptides from the spongin-based skeletons

of different demosponges. These studies show that the alkali

solution hydrolyses spongin. Obtained is a hydrolysate of

amino acids with no residual peptides visible on SDS-PAGE

gels after staining with Coomassie and the very sensitive

silver stain. These results agree well with those reported pre-

viously [22] about the strong insolubility of spongin, which

aimed to appropriate peptides for proteomics research.

Our recent findings of chitin within skeletons of both

marine demosponges from the Order Verongida [12,13] as

well as of hexactinellids [23] relied on the fact that spongin,

in contrast to chitin, is soluble in a 2.5 M NaOH solution.

Therefore, we used this simple test in the present study. We

decided to use the endemic freshwater sponge Lubomirskiabaicalensis (Pallas, 1773) for these investigations (figure 1a), as

the grey or brownish coloured holdfast of this sponge is par-

ticularly visible after it has been detached from stones or

other rocky substrates (figures 1b and 2a; electronic sup-

plementary material, figure S1). Initial experiments showed

dissolution of the holdfast-containing skeletal fragments after

2–4 h in 2.5 M NaOH at 378C. The presence of residual fibrous

matter could also be seen which strongly resembled the shape

of the sponge holdfast (figure 2; electronic supplementary

material, figures S3 and S4). Isolation of the holdfast in the

form of an alkali-resistant fibrous material still containing silic-

eous spicules motivated us to carry out, to our knowledge, the

first ever detailed analytical, biochemical and genetic investi-

gations to identify chitin as a possible candidate as the main

structural component of the sponge holdfast.

2. Material and methods(a) Sponge samplesSpecimens of L. baicalensis were collected in Lake Baikal near Bolshie

Koty Settlement (518540 1200 N, 1058060 0200 E) from 15–25 m depths

(water temperature 3–48C) by SCUBA during 2009–2012.

The samples collected were placed immediately in containers

with Baikal Lake water and ice, and transported to the

Limnological Institute SB RAS (Irkutsk) for 1.2 h at a constant

water temperature (3–48C).

(b) Isolation of chitin-based holdfastThe specimens of L. baicalensis were initially carefully inspected for

the intactness of their skeletons, and the presence of macroalgae or

invertebrates, using a stereomicroscope. Neither contaminants

nor damage were observed for the collected species. The isolation

of the chitin-based holdfast was performed according to the

alkali-based treatment steps as described in the electronic

supplementary material in details.

(a) (b)

Figure 1. (a) Underwater image of the endemic Baikal Lake sponge Lubomirskia baicalensis shows that this branched 50 cm tall demosponge is attached to therocky substrate. The bright green colour is due to a symbiotic algae (Zoochlorella) that lives in the external tissue layer of the sponge. (b) The sponges are attachedto the hard substrate via plate-like holdfast (arrows) that morphologically differ from fibrous spongin-based and silica spicules-containing skeleton. (Online versionin colour.)

(a) (b)

(c) (d)

Figure 2. The holdfast of L. baicalensis (a, arrow) became brownish during drying in air. The microstructure of the holdfast is quite visible using light microscopy(b – d ). The holdfast after 12 h of alkali treatment still shows light pigmentation and contains both spicules (b) and residual microparticles from the rocky substrateto that the sponge was attached (c). The alkali-resistant fibrous network within the holdfast becomes visible in the light microscope after 7 days of insertion in 2.5 MNaOH solution at 378C. Scale bars, 100 mm. (Online version in colour.)

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(c) Analytical methodsAnalytical methods like Raman spectroscopy, near-edge X-ray

absorption fine structure (NEXAFS) spectroscopy, Calcofluor

White (CFW) staining, as well as electrospray ionization

mass spectrometry (ESI-MS) and estimation of N-acetyl-D-gluco-

samine (NAG) contents are represented in the electronic

supplementary material.

(d) Chitin synthase gene detection from the genome ofLubomirskia baicalensis

Specimens of L. baicalensis to be used for RNA isolation were frozen

in liquid nitrogen; those to be used for DNA isolation were stored in

70 per cent ethanol at 48C. Total genomic DNA from sponge tissue

was extracted using the PureLink Genomic DNA kit (Invitrogen).

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Total RNAwas isolated from fresh or deep-frozen sponge specimens

using a Trizol Reagent kit (Sigma). cDNA was synthesized using

a Reverta kit (AmpliSens, Russia). Comparison of the known

chitin synthase mRNA sequences of freshwater sponge Spongillalacustris (HQ668146; HQ668147) and marine sponge Amphimedonqueenslandica (XP_003385441) revealed highly conserved regions

which have been chosen for designing several degenerate primers

(see the electronic supplementary material). PCR products obtained

with the primer pair of ChsFW_L1 (50-GGACATGTTGGATTCTG

ATCCCC-30) and ChsFW_R4 (50-CTCCGTGGATCAGGCAGC

TGAACTC-30) was subsequently cloned and sequenced (see the

electronic supplementary material).

Chitin synthase (CHS) genes were identified by compari-

son with the CHS sequences registered in GenBank using

‘BLAST-X’ tools at the National Center for Biotechnology

Information (NCBI) web site (http://www.ncbi.nlm.nih.gov).

The amino acid sequence encoded by the obtained CHS

cDNA was deduced using the software EDITSEQ (DNAStar Inc,

USA). The sequences obtained in this study were submitted

to GenBank and can be retrieved under the accession nos

JX875071–JX875074.

3. Results(a) Structural peculiarities of the Lubomirskia

baicalensis holdfastHabitat conditions of Baikal sponges differ considerably from

those of other freshwater sponges owing to hydrological and

hydrochemical peculiarities of Lake Baikal, such as great

depths, long ice periods, low water temperatures in summer

(10–128C) in the upper layers, high oxygen content and low

concentrations of organic matter [24]. Lubomirskia baicalensis(figure 1) has a branched shape and an encrusting base with

erect (30–60 cm up to 1 m high) dichotomous branches with

rounded apices. The diameter of branches varies from 1 to

4 cm ranging from cylindrical to flattened shapes. The colour

of live specimens is brilliant green, which is due to the

symbionts inhabiting the external layer of sponges [25].

Ectosomal skeleton consists of spicule tufts from primary spon-

gin fibres. The spicule skeleton consists of megascleres oxeas,

uniformly spined (145–233 � 9–18 mm; [26]). Lubomirskiabaicalensis is common on rocks, boulders and wood along the

entire shoreline at a depth from 3–4 m to more than 50 m.

Sponges could be easily mechanically detached from the

hard substrates such that the holdfast remained strongly

attached to the sponge body (figures 1b and 2a; see also elec-

tronic supplementary material, figure S1). Initially, we used

insertion of selected L. baicalensis specimens into 2.5 M

NaOH at 378C to examine the chemical stability of the

sponge skeletons to alkali treatment, because it is a well

known fact that spongin can be easily dissolved in solutions

that are up to 5 per cent alkali even at room temperature [21].

This stands in contrast to chitin, which is resistant to similar

alkali treatment at temperatures of up to 508C [12,13,23,27].

The light microscopy image (figure 2b) of L. baicalensis hold-

fast after 12 h incubation in alkaline solution shows the presence

of residual siliceous spicules as well as brownish pigmented

organic matter with some mineral microparticles (figure 2c)

that are resistant to the treatment. This organic material remains

undissolved in 2.5 M NaOH, even after incubation over 7 days

at 378C, and shows very intense characteristic fluorescence of

chitin after specific CFW staining (see the electronic supple-

mentary material, figure S2). Siliceous spicules, however, are

not more visible after this treatment in contrast to some mineral

particles. We observed that the alkali-resistant mineral micro-

particles of the substrate origin are still tightly bound into

chitinous fibres (figure 2c; electronic supplementary material,

figure S2). Because of this strong incrustation of the chitinous

fibrillar network with the mineral phase, we assume that

chitin and not spongin, which was dissolved during insertion

of the holdfast into alkaline solution, may be responsible for

attachment of the sponge to the rocky substrate.

The treatment of alkali-resistant matrix of the holdfast

with 3 M hydrochloric acid (HCl) leads to disappearance of

the mineral particles, however the fibrous matrix remains

stable. In our previously published work, we provided

strong evidence that chitin is also resistant to dissolution in

HCl [28]. This property can be also effectively used for iso-

lation of chitin that contains calcium carbonate-based

minerals as residual material. To confirm this discovery of

the presence of chitin within the holdfast of L. baicalensis,

we used a multitude of sensitive bioanalytical methods as

presented below.

(b) Identification of chitin within holdfast ofLubomirskia baicalensis

Recently, NEXAFS technique has been successfully applied to

determine key differences between electronic properties

related to the light adsorption by polysaccharides and pro-

teins, even within diverse biominerals [28–30]. We used

NEXAFS spectroscopy to explore site-specific electronic prop-

erties of L. baicalensis cleaned holdfast samples (measured on

500 � 500 mm areas) in order to gain insight into the nature of

the organic components. NEXAFS experiments, performed at

the carbon K edge, provided evidence that the contribution

of carbon is mainly owing to the organic part of the holdfast

(figure 3). Moreover, the carbon K-edge spectrum of this

sponge holdfast showed all the typical absorption features

of chitin and not those of spongin (figure 3), or collagen,

as was the case for spicules of the hexactinellid Hyalonemasieboldi in previous studies by our team [30]. Both chitin

and collagen spectra exhibited a strong peak at approxi-

mately 288 eV that is associated with the C 1s! p*

resonance involving acetamido (2NH(C ¼ O)CH3) group—

and pepty (–NH–C(O)–) group—character orbitals. Careful

inspection of the spectra indicates, however, that energies

of this peak are different for chitin (approx. 288.5 eV), spongin

(approx. 288.1 eV) and collagen (approx. 288.2 eV). It has been

shown [31,32] that the observed 0.3 eV shift manifests upon the

conversion of carboxyl bonds in lone amino acids into amide

bonds in peptide chains. While the 0.3 eV shift is rather

small, it has been well documented in the studies cited

above. It appears to provide a ‘sensible’ base for ‘in situ’ identi-

fication of polysaccharides and proteins—including naturally

occurring biocomposites, such as sponge holdfast—without

the need of preliminary disruption or extraction. In these ana-

lyses, we also show that the C¼O-character absorption peak of

the holdfast chitin is distinguishable from a strong cellulose

peak reported at 289.5 eV (see figure 3; [33]).

The chitin molecule consists of NAG (GlcNAc) residues,

including the acetamide group at the C-2 position of glucosa-

mine, the secondary hydroxyl group at C-3 and the primary

hydroxyl group at C-6 positions [34]. Therefore, estimation of

GlcNAc is the crucial step for chitin identification in organic

matrices of unknown origin.

0.8

0.6

0.4

0.2

2840

286 288 290E (eV)

285.0 eV

TE

Y (

arb.

uni

ts)

288.1 eV

288.52 eV

289.3 eV spongin skeleton

chitin

holdfast

cellulose(plant)

cellulose(bact.)

292 294

Figure 3. Detailed NEXAFS spectra taken at the C 1s threshold for L. baicalensiscleaned holdfast, spongin-based skeleton, chitin standard (Fluka) and two differ-ent celluloses of the plant and bacterial origin, respectively. Careful analysisreveals an energy shift approximately 0.3 eV of the C 1s! p* acetamido( – NH(C¼O)CH3) group peak in holdfast chitin relative to pepty ( – NH –C(O) – ) group position in collagen-like spongin. Furthermore, the spectra ofthe holdfast differ from those obtained for cellulose samples. (Online versionin colour.)

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Mass spectroscopy is one of the most sensitive methods

for analysis of D-glucosamine, which is the only product of

chitin acid hydrolysis. The obtained ESI-MS spectrum of

the hydrolyzed L. baicalensis holdfast sample is very similar

to the spectra of a D-glucosamine standard (see the electronic

supplementary material, figure S5), and consists of three

main signals with m/z ¼ 162.18, 180.02 and 359.61. The sig-

nals at m/z ¼ 180.02 clearly shows the presence of dGlcN

molecules in the sample and corresponds to a [M þ Hþ]

species of dGlcN (calculated molecular weight of 179.1).

The signal at m/z ¼ 162.06 corresponds to [M 2 H2O þ Hþ]

dGlcN ion (calculated: 162.1) which is the loss of one water

molecule [35,36]. The weak signal at m/z ¼ 359.13 corre-

sponds to [2 M þ Hþ] species which is the proton-bound

dGlcN non covalent dimer [36]. The sample at m/z ¼ 201 cor-

responds to [M 2 H2O þ Kþ] and [(GlcN)2 þ Kþ] adducts

with a potassium ion which is common for natural samples.

Interestingly, the sample can be completely hydrolyzed at

608C, but is stable in 6 M HCl at room temperature for

at least 24 h.

To quantify chitin in our samples, we measured the

amount of N-acethylglucosamine released by chitinases

using a Morgan–Elson colorimetric assay [37], which is the

most reliable method for the identification of alkali-insoluble

chitin owing to its specificity [38]. We detected 775.3+ 0.3 mg

N-acetyl-glucosamine per mg of L. baicalensis holdfast.

The results of Raman spectroscopy of the cleaned

L. baicalensis holdfast are represented in the electronic sup-

plementary material, figure S6. The Raman spectra for a-

chitin and L. baicalensis cleaned fibrous holdfast fibre material

were nearly identical, also in agreement with published reports

on chitin identification using Raman spectroscopy [39].

In summary, the analytical investigations described above

clearly show the presence of chitin within the holdfast of

L. baicalensis. This observation raises the question of the

presence of the corresponding CHS genes, which have thus

far not been described. We have, therefore, carried out the

corresponding genetic analysis, as described below.

(c) Characterization of chitin synthase genes inLubomirskia baicalensis

We have isolated and characterized four new CHS gene

fragments from the freshwater sponge L. baicalensis: CHS_LB01

(1088 bp), CHS_LB02 (924 bp), CHS_LB03 (1077 bp) and

CHS_LB04 (1232 bp). The first three sequences were identified

from sponge RNA, while CHS_LB04—from sponge DNA.

These sequences included the intron in position 935–1073

(139 bp). The sizes of the deduced hypothetical protein

fragments of Baikalian sponge CHSs were 308–364 aa. A com-

parison of CHS amino acid sequences yielded an identity of

74.5–92.5 per cent. BLAST-X analyses indicated that the pro-

tein sequences of CHS from Baikalian sponge were most similar

to CHS of S. lacustris, AEI55440 (73–98%), A. queenslandica,

XP_003385441 (52–54%), Hydra magnipapillata, XP_002162504

(39–43%), Nematostella vectensis, XP_001633545 (43–46%) and

Branchiostoma floridae, XP_002592717 (35–39%). CLUSTALX align-

ment of L. baicalensis CHS predicted proteins with those from

the closest relatives revealed the conserved domain specific

for all types of CHSs. This domain includes catalytically criti-

cal sequences GEDR and QRRRW [40] in the all amino acid

sequences (figure 4).

4. DiscussionThe presence of chitin in both evolutionary older marine and

evolutionary younger freshwater sponges [42] suggests that

the CHS genes found in this study represent shared ancestral

character states of sponges, and maybe even of a possible

common ancestor within the metazoan lineage (see the elec-

tronic supplementary material, figure S7). Thus, we suggest

that the chemistry, structure, morphology and biomechanical

properties of poriferan holdfasts were crucial throughout the

evolutionary history of sponges. Since dislodgment is

mostly fatal for adult sponges, the role of the holdfast is a critical

one. For sponges, holdfast morphology and sediment cohesive-

ness are important determinants of the maximum tensile

force they are able to withstand under specific environmental

conditions.

There are no doubts that fixation is very important for all

sedimentary animals, especially those which have little

opportunity to move or to re-build these body parts. Intrigu-

ingly, to our best knowledge, there are no reports regarding

the chitinous origin of the holdfast of other aquatic invert-

ebrates with exception of hydrorhiza of the hydroid

Myriothela cocksi [43]. Hydrorhiza is a rootstock by which a

hydroid is attached to other objects. The adhesion of the

hydrorhiza to the substratum is affected only by the perisarc

layer covering the flattened extremities of the adhesive tenta-

cles. This perisarc is about 6 mm thick and is composed of

true chitin [43].

The comparably smaller sponge-class Calcarea is charac-

terized by skeletal spicules of calcite, and holdfasts within

this taxon are less expressive, than those found among mem-

bers of the class Demospongiae, which are notably more

diverse, reach large sizes and inhabit more different solid

substrata (for review see [8]). Searching for specific types of

fixation including some hypothetical adhesive substances

for fixation in different classes of Porifera is a challenging

task for future studies.

Figure 4. Alignment of the C-terminal end of L. baicalensis CHS predicted proteins (L_baic_01, L_baic_02, L_baic_03 and L_baic_04) with predicted proteins fromSpongilla lacustris (S_lac_5605, GenBank accession no. AEI55440), Amphimedon queenslandica (Amph_queen, GenBank accession no. XP_003385441), Hydra magnipapillata(Hydra_magn, GenBank accession no. XP_002162504), Nematostella vectensis (Nemat_vect, GenBank accession no. XP_001633545) and Branchiostoma floridae (Bran_flori,GenBank accession no. XP_002592717). The alignment was performed using the CLUSTALX v. 2.0.10 program [41]. Amino acids that are conserved among eight to nine (whitesymbols on black background) and five to seven (black symbols on grey background) sequences are highlighted. The conservative domains (GEDR and QRRRW) of CHS aremarked with asterisks (*).

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Within the phylum Porifera, different attachment

methods have evolved over time. For example, the settled

larvae of Halichondria moorei undergoing metamorphosis

were found to possess a complex glycocalyx lining the cells

on their upper surface [44]. This structure, which has been

referred to as the sponge larval coat, was present on neither

adult sponges, nor on unsettled larvae. It was suggested

that this sponge’s mechanism for larval attachment bears

some similarity to the adhesion of many cultured cells to

their substrates. This hypothesis is supported by the absence

in sponge larvae of specialized cement glands, which are

known to be involved in substrate attachment in other

marine invertebrates [44].

Our finding of the presence of chitin within holdfast of

L. baicalensis suggests the existence of corresponding chitin-

producing cells. Intriguingly, these cells are definitively not

located within chitin-containing gemmules. In numerous fresh-

water sponges, gemmules consist of a rough coating (mainly

proteinaceous spongin and silicious gemmuloscleres) and

are filled with archaeocytic totipotent cells and many undifferen-

tiated and polynucleated cells [45]. It was found that newly

formed gemmular cells are first surrounded by a squamous cell

layer which later degenerates and becomes surrounded in turn

by a layer of columnar cells. The columnar cells secrete both

internal and external chitinous membranes. Jeuniaux [46]

reported that 1–3% of the dry weight of the gemmules’ coating

is chitin. However, while the production of dormant, encapsu-

lated gemmules by freshwater sponges and a few marine

species is a well-documented biological phenomenon [47],

there are to date no reports about gemmule production by any

species of the endemic family Lubomirskiidae, freshwater

sponges in the Baikal Lake, which is monophyletic with the

globally distributed Spongillidae [48]. Thus, an obvious object

of investigation with respect to identification of chitin is the

poorly investigated larva of L. baicalensis.One of the most intriguing questions that shall be

addressed by further investigations is to decipher the role of

the holdfast in sponge attachment, and how holdfasts maintain

their adhesive prosperities underwater and in highly saline as

well as in freshwater conditions. Such information may have a

high potential for use in biomimetic fields, where it could

be exploited for the design of biocompatible adhesives with

tuneable physico-chemical properties.

Financial support by DFG (projects EH 394/1–1) is gratefullyacknowledged. Furthermore, this study was funded by the ErasmusMundus DAAD Programme 2011, RFBR no. 11–04–00323-a, no. 12–02–00088-a, G-RISC (German-Russian Interdisciplinary ScienceCenter) grant 2012 and the CryPhysConcept Programme. We cor-dially thank Prof. Eike Brunner for use of the facilities at Instituteof Bioanalytical Chemistry, TU Dresden.

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Soc

B280:20130339

Electronic Supplementary Materials

First Report on Chitinous Holdfast in Sponges (Porifera)

Hermann Ehrlich1*

, Oksana V. Kaluzhnaya2, Mikhail Tsurkan

3,

Alexander Ereskovsky4, Konstantin R. Tabachnick

5, Micha Ilan

6, Allison Stelling

7, Roberta

Galli7, Olga V. Petrova

8, Serguei V. Nekipelov

8, Victor N. Sivkov

8, Denis Vyalikh

9,

René Born10

, Thomas Behm1, Andre Ehrlich

11, Lubov I. Chernogor

2, Sergei Belikov

2, Dorte

Janussen12

, Vasilii V. Bazhenov1, Gert Wörheide

12, 13, 14*.

1 Institute of Experimental Physics, TU Bergakademie Freiberg, 09599 Freiberg,

Germany

2 Limnological Institute SB RAS, 664033 Irkutsk, Russia

3 Leibniz Institute of Polymer Research & Max Bergmann Centre for Biomaterials,

01062 Dresden, Germany

4 Mediterranean Institute of Biodiversity and Ecology, CNRS, IRD, Aix-Marseille

University, 13007 Marseille, France

5 P.P. Shirshov Institute of Oceanology, Russian Academy of Sciences, Moscow,

Russia

6 Department of Zoology, Tel Aviv University, Tel Aviv 69978, Israel

7 Carl-Gustav-Carus Klinikum, TU Dresden, 01069 Dresden, Germany

8 Department of Mathematics Komi SC UrD RAS, Syktyvkar, Russia

9 Institute of Solid State Physics, TU Dresden, 01062 Dresden, Germany

10 Institute of Materials Science, TU Dresden, 01062 Dresden, Germany

11 Institute of Mineralogy, TU Bergakademie Freiberg, 09599 Freiberg, Germany

12 Forschungsinstitut und Naturmuseum Senckenberg, 60325 Frankfurt am Main,

Germany

13 Department of Earth and Environmental Sciences, Palaeontology & Geobiology,

Ludwig-Maximilians-Universität München, Richard-Wagner-Str. 10, 80333 München,

Germany

14 GeoBio-Center, Ludwig-Maximilians-Universität München, Richard-Wagner-Str.

10, 80333 München, Germany

15 Bayerische Staatssammlung für Paläontologie und Geologie, Richard-Wagner-Str.

10, 80333 München, Germany

Methods and Materials

Isolation of chitin-based holdfast

The isolation of the chitin-based holdfast was performed according to the following treatment

steps:

Step 1: The air dried sponge specimens containing holdfasts were cut into pieces of 1–3 cm,

placed in distilled water and sonicated at room temperature for 15 minutes to remove possible

microdebris, sediment particles, and associated microinhabitants from the investigated

fragments. Afterwards, the samples were rinsed three times with distilled water.

Step 2: These samples were treated with 2.5 M NaOH at 37°C for 12 h to remove proteins and

pigments [1]. The siliceous spicules are still present after 12 h of this treatment (they can be

dissolved only after 72 h under these conditions). Treatment was followed by three rinses with

distilled water.

It should be noted that the NaOH concentration of 2.5 M corresponds to 4% v/v. This is well

below the critical concentration of 25–30% v/v where the transformation of β-chitin into α-

chitin starts to take place [2]. Therefore, the crystal structure of the chitin-based skeletons was

not influenced by the extraction procedure. A commercially available crab α-chitin sample

was purchased from FLUKA. Monomeric N-acetyl-D-glucosamine was purchased from

Sigma–Aldrich.

Step 3: HCl treatment of sponge holdfast.

To elucidate the nature of the alkali resistant holdfast components of L. baicalensis under acid

dissolution, we carried out prompt demineralization of residual calcium carbonate-containing

microparticles from the rock using 3 M HCl. Alkali treated holdfasts were washed five times

in distilled water, and placed in a 10 mL glass vessel containing 7 mL of 3 M HCl solution.

The vessel was covered to restrict evaporation and was incubated at room temperature for

24 h. Immersion in HCl lead to an immediate loss of carbonate-based minerals from the acid

resistant fibrous organic matrix. The colorless material obtained after HCl-based treatment of

the sponge holdfast samples was washed with distilled water five times and brought up to a

pH level of 6.5, and finally dialyzed against deionized water on Roth (Germany) membranes

with a MWCO of 14 kDa. Dialysis was performed for 48 h at 4 °C. The dialyzed material was

dried at room temperature and used for staining and the different analytical investigations as

described below. The effects of the NaOH as well as of HCl treatment on the holdfasts of

L. baicalensis were examined using optical and fluorescence microscopy. Light and

fluorescence microscopy images were obtained with digital microscope Keyence BZ-8000.

Raman spectroscopy

Raman spectra were recorded on following apparatuses:

a) Kaiser HoloLab Series 2000 microscope coupled to an f/1.8 Holospec spectrograph (Kaiser

Optical Systems, Ann Arbor MI, USA) with a liquid nitrogen cooled CCD camera (Roper

Scientific, Trenton NJ, USA). A Toptica XTRA laser (Toptica Photonics AG, Gräfelfing,

Germany) with an excitation wavelength of 785 nm was used. The spectra were baseline-

corrected using a multi-point linear baseline (at 350, 800, 1700, 1960, 2600, 2750, and

3050 cm-1

).

b) Kaiser HoloLab Series 5000 microscope coupled to an f/1.8 Holospec spectrograph (Kaiser

Optical Systems, Ann Arbor MI, USA) with a liquid nitrogen cooled CCD camera (Roper

Scientific, Trenton NJ, USA). A Toptica XTRA laser (Toptica Photonics AG, Graefelfing,

Germany) with an excitation wavelength of 785 nm.

c) Bruker RFS 100/s spectrometer and Nd-YAG excitation at 1064 nm.

NEXAFS spectroscopy

The electronic structure of the L. baicalensis holdfast-free spongin-base, as well as the

holdfasts after the cleaning procedure described above, were characterized with near-edge X-

ray absorption fine structure (NEXAFS) spectroscopy at the Berliner Elektronenspeicherring

für Synchrotronstrahlung (BESSY) using radiation from the Russian-German dipole beamline

[3]. The cellulose of plant origin (Sigma-Aldrich, CAS Number 9004-34-6, EC Number 232-

674-9), bacterial cellulose (XCell, Xylos Corporation, Langhorne, PA, USA) and α-chitin

standard (Fluka) were used as references. All near-edge X-ray absorption fine structures were

acquired in the total electron yield (TEY) mode. The energy calibrations of the NEXAFS C-K

edge spectra have been performed by using the well resolved π-resonance at 285.38 eV of the

C-K edge spectrum of highly ordered pyrolytic graphit (HOPG) [4]. At that power the energy

resolution was less than 0.1 eV. To allow normalization of the incident radiation intensity, the

flux curve of the beamline was recorded using an Au photo cathode.

Calcofluor White Staining

To elucidate the particular location of chitin in the skeletal structures, we used Calcofluor

White (Fluorescent Brightener M2R, Sigma), which shows enhanced fluorescence when

binding to chitin [5,6]. Pieces of untreated clean skeleton holdfasts, and those dialyzed after

alkali and HCl treatment, were placed in 0.1M Tris-HCl at pH 8.5 for 30 min. Then, they

were stained using 0.1% Calcofluor White solution for 30 min in darkness, and rinsed three

times with distilled water. They were next dried at room temperature, and finally observed

using fluorescence microscopy. It should be noted here that CFW-stained chitin is highly

visible due to the strong blue fluorescence, even at exposure times between 1/50 and 1/500

seconds.

Electrospray ionization mass spectrometry ESI-MS

Organic matrix obtained after NaOH and HCl-treatment of L. baicalensis holdfasts was

hydrolyzed in 6 M HCl for 24 hours at 60 °C. Solid remains were filtered from the sample

with a 0.4 micron filter and freeze-dried to remove the excess HCl. The solid remains were

dissolved in a water-acetonitrile mixture for ESI-MS analysis. The standard D-glucosamine

was purchased from Sigma (USA) and its spectrum was measured under similar conditions.

All ESI-MS measurements were performed on Waters TQ Detector ACQUITYuplc mass

spectrometer (Waters, USA) equipped with ACQUITYuplc pump (Waters, USA) and

BEHC18 1.7 mm 2.1x50mm UPLC column. Nitrogen was used as the nebulizing and

dessolvation gas. Graphs were generated using Origin 8.5 for PC

Estimation of N-acetyl-D-glucosamine (NAG) contents

Preparation of colloidal chitin from a crab α-chitin standard (Sigma) was performed according

to [7]. The Morgan-Elson assay was used to quantify the N-acetyl-D-glucosamine released

after chitinase treatment as described previously [7].

Purified and dried L. baicalensis sponge holdfast samples (6 mg) were pulverized to a fine

powder in an agate mortar. The samples were suspended in 400 ml of 0.2 M phosphate buffer

at pH 6.5. A positive control was prepared by solubilizing 0.3% colloidal chitin [5] in the

same buffer. Equal amounts of 1 mg/mL of three chitinases (EC 3.2.1.14 and EC 3.2.1.30): N-

acetyl-D-glucosaminidase from Trichoderma viride (Sigma, No. C-8241), and two poly (1,4-

α-[2-acetamido-2-deoxy-D-glucoside]) glycanohydrolases from Serratia marcescens (Sigma,

No. C-7809), and Streptomyces griseus (Sigma, No. C-6137) were suspended in 100 mM

sodium phosphate buffer at pH 6.0.

Digestion was initiated by mixing 400 ml of the sample and 400 ml of the chitinase mix.

Incubation was performed at 37° C and stopped after 114 h by adding 400 ml of 1% NaOH,

followed by boiling for 5 min. The vessels were centrifuged at 7000 rpm for 5 min and the

purified reducing sugars were used for a 3, 5-dinitrosalicylic acid assay (DNS) [8]. For this

purpose, 250 ml of the supernatants and 250 ml of 1% DNS were dissolved in a solution

containing 30% sodium potassium tartrate in 0.4M NaOH. The reagents were mixed and

incubated for 5 min in a boiling water bath. Thereafter, the absorbance at 540 nm was

recorded using a Tecan Spectrafluor Plus Instrument (Mannedorf/Zurich, Switzerland). Data

were interpolated using a standard curve prepared with a series of dilutions (0–3.0 mM) of N-

acetyl-D-glucosamine (Sigma, No. A-8625) and DNS. A sample, which contained chitinase

solution without substrate, was used as a control.

Chitin-synthase gene detection from the genome of L. baicalensis

Comparison of the known chitin-synthase mRNA sequences of freshwater sponge Spongilla

lacustris (HQ668146; HQ668147) and marine sponge Amphimedon queenslandica

(XP_003385441) revealed highly conserved regions which have been chosen for designing

several degenerate primers. The primers were synthesized by Eurogen (Russia):

ChsFW_L1: 5’-GGACATGTTGGATTCTGATCCCC-3’

ChsFW_R1: 5’-GGGGATCAGAATCCAACATGTCC-3’

ChsFW_L2: 5’-GACATGGGTGAAGACCGCTGGCTGTGCAC-3’

ChsFW_R2: 5’-GTGCACAGCCAGCGGTCTTCACCCATGTC-3’

ChsFW_L3: 5’-CTATTCACTCTGCTGTTCATC-3’

ChsFW_R3: 5’-GATGAACAGCAGAGTGAATAG-3’

ChsFW_L4: 5’-GAGTTCAGCTGCCTGATCCACGGAG-3’

ChsFW_R4: 5’-CTCCGTGGATCAGGCAGCTGAACTC-3’

Primer pairs were checked with different combinations and the pair ChsFW_L1 and

ChsFW_R4 gave the positive PCR-signal and was selected for future experiments. PCR

amplification of chtin-synthase gene fragment was performed on a Peltier Thermal Cycler

(MJ Research, USA) using the Taq PCR Master Mix Kit (QIAGEN, Germany). Cycling

conditions were follows: initial denaturation at 95°C for 5 min, 35 cycles of 95°C for 40 s,

57°C for 1 min, and 72°C for 1 min 30 s, and a final extension of 10 min at 72°C. Amplicons

were separated by 1% agarose gel electrophoresis. PCR products of expected size (1000-

1200 bp) was recovered from gel using PCR Clean-Up Gel Extraction NucleoSpin Extract II

kit as recommended by Macherey-Nagel (Germany), ligated into the pTZ57R/T (Fermentas)

vector and transformed into CaCl2-competent Escherichia coli XL1BL. Positive clones were

identified by PCR amplification with universal M13-20 (5’-GTA AAA CGA CGG CCA GT-

3’) and M13-reverse (5’-CAG GAA ACA GCT ATG AC-3’) primers (Evrogen, Russia) using

following cycling conditions: initial denaturation at 95°C for 2 min, 35 cycles of 95°C for

20 s, 58°C for 40 s, and 72°C for 1 min 20 s. The plasmid DNAs from 35 arbitrary selected

clones were extracted with the QIAGEN plasmid kit (QIAGEN, California, USA).

Sequencing was performed using M13-20 and M13-reverse primers on CEQ 8800 Genetic

Analysis System using DTSC Quick Start Kit (Beckman Coulter, USA). Sequence data were

edited by BioEdit and the amino acid sequence encoding by the obtained CHS cDNA was

deduced using the software EditSeq (DNAStar Inc, USA).

The amino acid sequences of chs genes available in GenBank were used to make a

phylogenetic analysis. Phylogenetic reconstructions were performed with maximum

likelihood (ML) approaches using the Molecular Evolutionary Genetics Analysis software

(MEGA v5.1.) [9]. The robustness of ML tree topologies was evaluated by bootstrap analysis

based on 1000 replicates. The most appropriate evolutionary model of amino-acid substitution

was defined as WAG+G. On phylogenetic tree (ESM Fig. 6) the chs genes from Fungi and

Insecta formed independent clades with high bootstrap-values. Chitin synthases from

freshwater and marine sponges formed separated branch within the group which also includes

phylum Chordata, Cnidaria and Choanoflagellata.

Supplementary figures:

Supplermentary figure 1. The holdfasts are well visible on the basal parts of Lubomirskia

baicalensis (a). They can be simply cut off from the sponge body (b) of the freshly collected

(b, d) as well as dried specimens from museum collections (c).

Supplementary figure 2. The alkali treated (7 days at 37°C) fibrous network of the

L. baicalensis holdfast (a) becomes more clearly visible in the fluorescence microscope (b)

after specific Calcofluor White staining for chitin identification. The alkali resistant mineral

microparticles of the substrate origin (arrows) are tightly bound to chitinous fibers. (Light

exposure time ½ s, scale bars: 100 µm).

Supplementary figure 3. Confocal microscopy image of the naturally occurring holdfast of

L. baicalensis prior to alkali treatment shows brownish colored fibers of spongin (white

arrows) tightly bond into chitinous template (scale bar: 100 µm).

Supplementary figure 4. Confocal microscopy image of the holdfast of L. baicalensis during

alkali treatment shows disappearance of the spongin fibers (black arrows). The chitinous parts

of the holdfast including mineral particles remain intact (scale bar: 100 µm).

150 200 250 300 350 400 450 500

0.0

2.0x107

4.0x107

6.0x107

150 200 250 300

0.0

3.0x107

6.0x107

9.0x107

a.i.

Mw/z

standard180.07

162.06

a.i

.

Mw/z

sample

162.18

180.02

201.7359.67

380.41

Supplementary figure 5. Electrospray ionization mass spectrum of the hydrolysed

L. baicalensis holdfast sample with inclusion of the ESI-MS spectra of D-glucosamine

standard.

Supplementary Figure 6. Raman spectra of purified chitin isolated from the holdfast of

L. baicalensis (a) shows high similarity to the Raman spectra of the α-chitin standard (b).

Supplementary Figure 7. Phylogenetic analysis (ML) of chitin synthase gene fragments

from different organisms. Sequences obtained from L. baicalensis are marked with dark

circles and written in bold. The chs genes from Fungi, Ecdysozoa and Amoebozoa formed

independent clade with high bootstrap-values. Chitin synthases from the freshwater sponge

L. baicalensis belong to a clade that includes Choanoflagellata, Porifera, Cnidaria, and

Chordata. Here, chs gene fragments from L. baicalensis form clade with those from the

freshwater sponge S. lacustris (marked with white circles) and marine sponge Amphimedon

queenslandica.

1855,4 1600 1400 1200 1000 800 600 434,4

1,0

10

20

30

40

50

60

70

80

90

99,2

cm-1

%T

Supplementary Figure 8. FTIR spectra of α-chitin standard (Fluka) (gray line) are nearly

identical to chitin isolated from the holdfast of L. baicalensis (brown line) as well as to chitin

from Aplysina aerophoba marine sponge (blue line).

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