and polyol dehydrogenases in patella vulgata and · grupos de organismos. ... o segundo maior filo...

Aromatic alcoholand polyoldehydrogenases inPatella vulgata andother gastropods(Mollusca)Diogo Ribeiro Amaral de CarvalhoMestrado em BioquímicaDepartamento de Química e Bioquímica

2015/2016

Orientador

Alexandre Lobo da Cunha, Professor Catedrático, Instituto de

Ciências Biomédicas Abel Salazar

Coorientador

Vitor Costa, Professor Associado, Instituto de Ciências Biomédicas

Abel Salazar

Todas as correções determinadas

pelo júri, e só essas, foram efetuadas.

O Presidente do Júri,

Porto, /_ /

FCUP/ICBAS/i3S Aromatic alcohol and polyol dehydrogenases in Patella vulgata and other gastropods (Mollusca)

I

Agradecimentos

Aos meus orientadores: ao Professor Doutor Alexandre Lobo-da-Cunha, pelo

apoio, interesse e auxílio prestados na orientação deste trabalho ao longo destes quase

cinco anos de investigação conjunta; e ao Professor Doutor Vitor Costa, por ter aceite o

desafio de auxiliar a investigação de um organismo tão estranho e muito pouco

conhecido pela comunidade científica em geral.

À equipa técnica do laboratório de Biologia Celular do departamento de

Microscopia do ICBAS, com especial destaque para a Ana Paula Sousa, pelas

infindáveis e martirizantes quantidades de tubos de ensaio lavados em prol deste

trabalho.

À equipa técnica do departamento de Biologia Molecular do ICBAS,

nomeadamente à Carla Oliveira e à Marisa Castro, pela cedência de equipamento e

apoio prestado na realização de alguns procedimentos laboratoriais.

À equipa do grupo de investigação UP3 do IBMC/i3S, nomeadamente ao Doutor

Frederico Silva à Fátima Fonseca, pelo apoio prestado na realização dos procedimentos

cromatográficos.

Ao Doutor Hugo Osório do IPATIMUP/i3S, pelo apoio prestado na identificação e

sequenciação das proteínas das bandas dos géis.

Aos meus, sempre presentes, colegas de mestrado, que individualmente foram

partilhando comigo as suas vivências, familiarizando-me com o que significa ser

bioquímico; e não esquecendo a origem de cada um, aceitam agora um novo conceito,

o de Bioquímico do Meio Aquático.

To you, Lucie, because you were a very important part of my life in this biochemical

journey, and in your six months’ adventure in Portugal, we were still able to share some

unforgettable moments together.

Ao pessoal do Centro Português de fotografia, à equipa de seguranças e às

minhas colegas, nomeadamente à Maria da Luz, a que desejo rápidas melhoras, e mais


II

recentemente à Mónica; um grande obrigado por tudo aquilo que me ensinaram nestes

três meses e meio de experiência como trabalhador-estudante.

And to you, my dear Alex, that like a tornado that comes from the sea to make

changes in land, so your overseas arrival testified the end of this chapter of my life, and

brought a new one alive; hopefully for many long years…


III

Resumo

As desidrogenases alcoólicas são enzimas bem conhecidas, estudadas em muitos

grupos de organismos. No entanto, pouco se sabe sobre estas enzimas em moluscos,

o segundo maior filo de animais multicelulares. Em estudos preliminares com várias

espécies dos cinco principais clados de gastrópodes, nomeadamente

Patellogastropoda, Neritimorpha, Vetigastropoda, Caenogastropoda e Heterobranchia,

foi possível detetar desidrogenases de álcoois aromáticos (usando álcool cinamílico

como substrato) e poliálcoois (usando manitol e sorbitol como substrato) em

homogeneizados totais de glândula digestiva. Usando a fração citosólica da glândula

digestiva de Patella vulgata, Phorcus lineatus, Littorina littorea e Siphonaria pectinata, a

atividade destas enzimas foi detetadas in situ após eletroforese em gel nativo de

poliacrilamida, verificando-se um perfil enzimático semelhante entre as várias espécies.

Este trabalho tinha como objetivo o isolamento e identificação destas enzimas, da

glândula digestiva da lapa Patella vulgata, uma espécie abundante na zona intertidal

portuguesa. Frações enriquecidas nas enzimas de interesse foram obtidas por

cromatografia de afinidade e de troca iónica. Estudos de proteómica revelaram a

presença de várias desidrogenases, onde se incluem uma álcool desidrogenase de

classe III e duas desidrogenases de poliálcoois (sorbitol e arabinitol desidrogenases) A

clonagem dos genes que codificam para estas enzimas e a expressão heteróloga destas

proteínas, irá permitir a sua caracterização, incluindo a atividade das desidrogenases

para diversos substratos alcoólicos.

Palavras-Chave

Desidrogenases alcoólicas, moluscos, gastrópodes, álcoois aromáticos,

poliálcoois, glândula digestiva, arabinitol desidrogenase, sorbitol desidrogenase.


IV

Abstract

Alcohol dehydrogenases are well known enzymes, studied in many types of

organisms. However, little is known about these enzymes in molluscs, the second largest

multicellular animal phylum. Preliminary studies with several species from the five major

gastropod clades, namely Patellogastropoda, Neritimorpha, Vetigastropoda,

Caenogastropoda e Heterobranchia, allowed the detection of aromatic alcohol

dehydrogenases (using cinnamyl alcohol as substrate) and polyalcohol dehydrogenases

(using mannitol and sorbitol as substrate) in digestive gland total homogenates. Using

the digestive gland cytosolic fraction of Patella vulgata, Phorcus lineatus, Littorina littorea

e Siphonaria pectinata, the activity of these enzymes was also detected in situ after

native polyacrylamide gel electrophoresis which resulted in a similar overall profile in all

studied species. This work aimed at the isolation and identification of these enzymes

from the digestive gland of the limpet Patella vulgata, an abundant species in the

Portuguese intertidal zone. Fractions enriched in the enzymes of interest were obtained

by affinity and ion exchange chromatography. Proteomic studies revealed the presence

of several dehydrogenases, which are included one class III alcohol dehydrogenase and

two dehydrogenases (sorbitol and arabinitol dehydrogenase). Cloning of the genes

encoding for these enzymes and the heterologous expression of these enzymes will

allow their characterization, including the dehydrogenase activity for diverse alcoholic

substrates.

.

Keywords

Alcohol dehydrogenases, molluscs, gastropod, aromatic alcohol, polyalcohol,

digestive gland, arabinitol dehydrogenase, sorbitol dehydrogenase.


V

Index

List of figures and tables .............................................................................................. VI

List of abbreviations ..................................................................................................... IX

1. Introduction ............................................................................................................ 10

1.1. Gastropod taxonomy and phylogeny ............................................................. 10

1.2. Digestive Gland ............................................................................................. 14

1.3. Oxidation of polyols and aromatic alcohols in gastropods ............................. 16

1.4. Alcohol dehydrogenases ............................................................................... 18

1.4.1. MDR - Type I ADHs ............................................................................. 18

1.4.2. SDR - Type II ADHs ............................................................................. 21

1.4.3. LDR - Type III ADHs ............................................................................ 22

1.4.4. Polyol Dehydrogenases ....................................................................... 23

1.5. Alcohol and polyol dehydrogenases in molluscs............................................ 26

1.6. Aims of this work ........................................................................................... 27

2. Materials and Methods ........................................................................................... 28

2.1. Species and sampling sites ........................................................................... 28

2.2. Preparation of protein extracts ...................................................................... 28

2.3. Enzyme activities .......................................................................................... 30

2.4. Affinity chromatography ................................................................................. 31

2.5. Ion exchange chromatography ...................................................................... 31

2.6. SDS-polyacrylamide gel electrophoresis ....................................................... 31

2.7. Protein sequencing ....................................................................................... 32

3. Results ................................................................................................................... 33

3.1. Enzymatic activities ....................................................................................... 33

3.2. Protein isolation and identification ................................................................. 36

4. Discussion .............................................................................................................. 45

5. Conclusions and perspectives ................................................................................ 49

6. References ............................................................................................................. 50


VI

List of figures and tables

Figure 1 - Major gastropod clades, according to morphological and molecular data ... 11

Figure 2 - Pictures of some gastropod species included in this study. A – Patella vulgata,

B – Phorcus lineatus, C – Nucella lapillus, D – Nassarius reticulatus, E – Siphonaria

pectinata, F – Armina maculata, G – Bulla striata, H – Aglaja tricolorata (provided

by A. Lobo-da-Cunha, University of Porto). .......................................................... 13

Figure 3 - Schematic representation of the digestive gland cells in gastropods. mv –

microvilli, ev – endocytic vesicle, ly – lysosome, li – lipid droplets, nu – nucleus, sv

– secretory vesicles, cc – calcium concretions (provided by A. Lobo-da-Cunha,

University of Porto). ............................................................................................. 15

Figure 4 - [A] Ultrastructure of the basophilic cells of the digestive of A. depilans, showing

tubular structures in the interior of rough endoplasmic reticulum cisternae (*) [12].

[B] Frozen section of the digestive gland of A. depilans showing stained basophilic

cells (arrows) after the detection of mannitol oxidase activity [12]. ....................... 16

Figure 5 - Scheme of the polyol pathway of the glucose metabolism. Conversion of

glucose to sorbitol by aldose reductase, and sorbitol to fructose by sorbitol

dehydrogenase [98]. ............................................................................................ 24

Figure 6 - Structure of sheep sorbitol dehydrogenase

(http://www.rcsb.org/pdb/explore/explore.do?structureId=3QE3). ........................ 24

Figure 7 - Native electrophoresis gel sections with in situ detection of the activity of

dehydrogenases of several alcoholic substrates, in four marine herbivorous species.

............................................................................................................................ 35

Figure 8 - Ion exchange chromatogram of 10 mM NAD+ fraction from affinity

chromatography with HiTrap DEAE Sepharose FF column. Proteins eluted with an

ascending and continuous NaCl gradient (0.05 M to 1 M), at a flow rate of 1 ml per

sec. (blue) protein detection (abs 280 nm); (green) high salt solution concentration;

(red) conductivity. ................................................................................................ 38

Figure 9 - SDS-polyacrylamide gel showing the enzymatic profile of some ion exchange

chromatography fractions obtained with HiTrap DEAE Sepharose FF column


VII

(Figure 8). Proteins identified by MS/MS using a database of Patella vulgata: 1 –

glutathione-S-transferase mu class; 2 – no statistical significant identification; 3 – D-

arabinitol dehydrogenase 1; 4 – cytosolic malate dehydrogenase; 5 – sorbitol

dehydrogenase-like protein (Table 7). .................................................................. 39

Figure 10 - Ion exchange chromatogram of 10 mM NAD+ fraction from affinity

chromatography with Mono Q 4.6/100 PE column Proteins eluted with an ascending

and continuous NaCl gradient (0.05 M to 1 M), at a flow rate of 1 ml per sec. (green)

protein quantification (abs 280 nm); (black) high salt solution concentration; (red)

conductivity. ......................................................................................................... 42

Figure 11 - Characterization of the fractions obtained by ion exchange chromatography

(Mono Q 4.6/100 PE). Top: analysis of polyol dehydrogenase activity. Bottom:

analysis of proteins by SDS-PAGE. Proteins were stained with Coomassie blue.

Molecular weight markers are shown on the left and right. Spots 1-7 were identified

by MS/MS: 1 & 2 alcohol dehydrogenase class III; 3 – universal stress protein; 4 &

5 - D-arabinitol dehydrogenase and Glyceraldehyde-3-Phosphate dehydrogenase

1; 6 & 7 – sorbitol dehydrogenase-like protein (Table 8). ..................................... 43

Table 1 - Compatible substrates for human liver sorbitol dehydrogenase [105] ........... 26

Table 2 - Taxonomy, habitat and diet of the 26 species of gastropods included in this

study. Ma - marine, Fw - freshwater, Te - terrestrial ............................................. 29

Table 3 - Activity of dehydrogenase in the digestive gland of gastropods. Values are

mean ± SD of at least 5 independent experiments (nmol.min-1. g-1 ± SD). Shaded

lines correspond to carnivorous species. ............................................................. 34

Table 4 - The cytosolic fraction of the digestive gland of Patella vulgata was loaded on a

Blue Sepharose CL-6B column, the flow-through was recovered and the column

washed with buffer. Proteins were eluted with NAD+ solutions of increasing

concentrations (5 mM, 10 mM and 20 mM). The column was then washed with

regeneration buffers. Dehydrogenase activities for polyols (D-mannitol and D-

sorbitol) and cinnamyl alcohol (aromatic) in the fractions were assayed by

spectrophotometry. A/ml – activity per ml of fraction; A/mg – activity per mg of

protein (specific activity); A total – activity of total fraction volume. ...................... 37


VIII

Table 5 - The cytosolic fraction of the digestive gland of Patella vulgata was loaded on a

Blue Sepharose CL-6B column, the flow-through was recovered and the column

washed with buffer. Proteins were eluted with two NAD+ solutions of different

concentrations (10 mM and 20 mM). The column was then washed with

regeneration buffers. Dehydrogenase activities for polyols (D-mannitol and D-

sorbitol) and cinnamyl alcohol (aromatic) in the fractions were assayed by

spectrophotometry. A/ml – activity per ml of fraction; A/mg – activity per mg of

protein (specific activity); A total – activity of total fraction volume. ...................... 37

Table 6 - 10 mM NAD+ fraction from affinity chromatography loaded on a HiTrap DEAE

Sepharose FF column. Proteins eluted with an ascending and continuous NaCl

gradient (0.05 M to 1 M), at a flow rate of 1 ml per sec. (Figure 8). Distribution of

dehydrogenase activities for polyols (D-mannitol and D-sorbitol) and cinnamyl

alcohol (aromatic) in some fractions were assayed by spectrophotometry. A/ml –

activity per ml of fraction; A/mg – activity per mg of protein (specific activity); A total

– activity of total fraction volume. Fractions: no.4 – flow-through representative, no.

22, 25 and 27 – protein peaks representatives. ................................................... 37

Table 7 - Proteins of interest (Figure 9) were analysed by mass spectrometry (MS/MS)

and identified using a Patella vulgata database. Blastp of the identified proteins was

performed at the NCBI database. Mascot search result scores greater than 65 are

significant (p<0.05). ............................................................................................. 40

Table 8 - Proteins from ion exchange chromatography fraction with Mono Q 4.6/100 PE

column (Figure 11) analysed by mass spectrometry (MS/MS) and identified with a

Patella vulgata database. Mascot search result scores greater than 65 are

significant (p<0.05). Data shows the highest score identifications for each sample.

............................................................................................................................ 44

Table 9 - Peptide ions from ion exchange chromatography fraction with Mono Q 4.6/100

PE column (Figure 11) analysed by mass spectrometry (MS/MS) and identified with

UniProt (Metazoa) database. Mascot search result scores greater than 82 are

significant (p<0.05); Data shows the highest score identifications for each sample.

............................................................................................................................ 44

Table S1 - Oxidase activities in the digestive gland of gastropods (nmol.min-1. g-1 ± SD).

Shaded lines correspond to carnivorous species (data not published). ................ 58


IX

List of abbreviations

RNA Ribonucleic Acid

NAD(H) Nicotinamide Adenine Dinucleotide

FAD Flavin Adenine Dinucleotide

ADH Alcohol Dehydrogenase

NADP(H) Nicotinamide Adenine Dinucleotide Phosphate (reduced form)

MDR Medium-chain Dehydrogenase/Reductase

DNA Deoxyribonucleic Acid

SDR Short-chain Dehydrogenase/Reductase

PGDH 15-Hydroxyprostalandin Dehydrogenase

LDR Long-chain Dehydrogenase/Reductase

HOT Hydroxyacid-Oxoacid Transhydrogenase

SDH Sorbitol Dehydrogenase

ALR Aldose Reductase

EDTA Ethylenediaminetetraacetic Acid

PMSF Phenylmethylsulfonyl Fluoride

NBT Nitro Blue Tetrazolium Chloride

TCA Trichloroacetic Acid

SDS Sodium Dodecyl Sulfate

MS Mass Spectrometry

MALDI Matrix-Assisted Laser Desorption Ionization

GAPDH Glyceraldehyde 3-phosphate Dehydrogenase

ARDH Arabinitol Dehydrogenase


10

1. Introduction

1.1. Gastropod taxonomy and phylogeny

Gastropoda is a class of molluscs that includes terrestrial, marine and freshwater

snails and slugs, and it was traditionally divided in three subclasses: Prosobranchia,

Opisthobranchia e Pulmonata [1]. However, morphological and molecular studies

showed that these were not monophyletic groups.

In the last decades, many studies have been dedicated to gastropod phylogeny

[2]. These studies rely mostly in molecular methods to establish phylogenetic trees and

verify group monophyly. Nevertheless, despite the increasing use of rRNA and

mitochondrial genomes, there are still a lot of doubts regarding phylogenetic

relationships between gastropod clades. Thus, gastropods are now divided in the clades:

Patellogastropoda, Neritimorpha, Cocculiniformia, Neomphalina, Vetigastropoda,

Caenogastropoda e Heterobranchia (Figure 1) [2-4].

The clade Patellogastropoda is composed by marine gastropods, known as true or

patellid limpets, which can be found attached to almost any rocky surface, coralline

algae, sunken vessels, whale bone and even to the shell of other molluscs. Their

distribution knows no borders, from the intertidal rocky shores to the deep-sea waters,

from polar to tropical region, and they play very important roles in littoral ecosystems,

having been largely used in ecological studies [5]. These are nocturnal grazing molluscs

that, by scraping the rock surface with their radulae ingest diatoms, settling stages of

algae and invertebrates, filamentous algae, cyanobacteria and detritus. This way, large

rocky areas are maintained in a primary stage of succession [6].

Among all Patellogastropoda families, Patellidae is the most studied one with a

total of 38 species in 4 genera., based in morphological studies. The genus Patella

comprises nine species, that occur in the north-eastern Atlantic area. This genus has

been used for many genetic studies, although very few species were analysed or they

represent a small region [5]. A very common limpet in the Portuguese coast is the species

Patella vulgata (Figure 2A). On the other hand, the family Lottiidae is the most diverse

within Patellogastropoda, comprising 130 species in at least 8 genera, with a worldwide

distribution of temperate and tropical areas. It includes the first mollusc species with full

sequenced genome (Lottia gigantea), being important as a representative of gastropod

basal lineages [5].


11

Figure 1 - Major gastropod clades, according to morphological and molecular data

Neritimorpha is a group of large morphological variety, such as the shell shape.

Besides the typical snail appearance, some species resemble limpets or slugs. They are

adapted to some habitats of harsh conditions, like deep-sea hydrothermal vents and

submarine caves, but also distributed in intertidal and subtidal areas of tropical regions,

freshwater habitats and arboreal terrestrial environments [7]. Most neritimorphs are

grazers on algal spores, diatoms and detritus [8]

Regarding recent molecular studies, the groups Neophalina and Cocculinoidea are

now considered two deep-water sister clades. Cocculinids is a group of small, white-

shelled deep-sea limpets, found in decaying squid beaks and sunken-bone/wood, while

neomphalines share features with both cocculinids and neritimorphs and are found

exclusively in hydrothermal vents, hydrocarbon seeps and sunken-wood [3]

The Vetigastropoda clade is constituted by marine snails, largely distributed across

all oceans, from deep waters to the shallow intertidal areas. They display a large variety

of shell morphologies, such as conical, limpet and trochiform types (Figure 2B), that can

also contain holes, slits and fissures [3]. They feed on encrusting invertebrates, however

Heterobranchia

Caenogastropoda

Vetigastropoda

Neritimorpha

Patellogastropoda

“Basal” Heterobranchia

Neomphalina

Cocculiniformia

Euthyneura Nudipleura

Tectipleura Euopisthobranchia

Panpulmonata


12

some groups, such as Trochoidea, can feed on algae and plant material such as marine

angiosperms [9].

Caenogastropoda is the largest and most diverse group of living snails, with about

136 existing families. It is a group of ecological and commercial important marine

families, but also contains some terrestrial and freshwater representatives. They present

a wide array of shell morphologies (coiled, uncoiled, globose, elongate, limpet-shaped)

(Figure 2C,D), however, some species present a reduced shell or more rarely do not

contain a shell. Caenogastropoda are found in a wide range of habitats, such as pelagic

drifters or active swimmers, benthic epifaunal or burrowers, herbivores, grazers, active

carnivores, detritus or sedentary suspension feeders, ectoparasites or shell-less

endoparasites [10].

Heterobranchia is a large clade formed by marine, freshwater and terrestrial snails

and slugs, which taxonomy have been changed mainly due to molecular data. This clade

is characterized by several apomorphic features [4], comprising several groups which

systematic position is not yet clear. Those groups presenting more ancestral features

are included in the informal group of “Basal heterobranchs”, which are clearly

paraphyletic [11]. The clade Euthyneura is characterized by a typical non/crossed

nervous system and innervated rhinophores. It is divided in Nudipleura, that occupy a

basal position, suggesting some plesiomorphic features despite the derived ones (Figure

2F), and Tectipleura that is characterized by a monaulic genital system. This last group

is divided in Euopisthobranchia, that share a cuticularized oesophagus (Figure 2G,H),

and Panpulmonata that presents a double-root procerebrum (Figure 2E). Overall, the

antique Euthyneura clade is the only who will survive to the molecular revolution of

gastropod systematics [4]. Regarding their dietary habits, heterobranchs can be

herbivorous, carnivorous, omnivorous or detritivorous, playing an active role hunting

pursuing their pray, or more sedentary one, scavenging or grazing the nearby territory

[11].


13

Figure 2 - Pictures of some gastropod species included in this study. A – Patella vulgata, B – Phorcus lineatus, C – Nucella

lapillus, D – Nassarius reticulatus, E – Siphonaria pectinata, F – Armina maculata, G – Bulla striata, H – Aglaja tricolorata

(provided by A. Lobo-da-Cunha, University of Porto).


14

1.2. Digestive Gland

The digestive gland is a major organ of the molluscs’ digestive system, being

composed by a vast number of tubules with a dead end, connected to the stomach via a

system of branched ducts. This organ is involved in extracellular and intracellular

digestion of food, nutrient absorption, lipid, glycogen and minerals storage, also having

an important role in detoxification [12-14].

The digestive tubules, also called digestive diverticula, have a one-cell layer

epithelium, surrounded by the basal lamina and connective tissue with muscular cells.

The pulsatile contractions of the muscular cells, combined with the stomach contractions,

move the food materials into the digestive gland. On the other hand, the beating of the

ciliary apparatus of the epithelial cells drives the materials away from the digestive gland,

toward the stomach and intestine. These seem to be the two systems, of opposite

directions that carry the ingested material through the ducts. Inside the ducts, fine

particles and soluble materials are taken up rapidly by endocytosis, however larger

particles are moved directly from the stomach to the intestine, being therefore excluded

from the digestive gland ducts [14].

The detoxification role of the digestive gland is inferred due to its capacity to

accumulate passively ingested metals, correlated with the occurrence of metal binding

proteins. The accumulation of metals can result in severe damage of the digestive gland

cells, when compared with other regions of the digestive tract [14].

In gastropods, the digestive gland epithelium is mainly formed by digestive and

basophilic cells (Figure 3), but in some species it may contain additional cell types [15,

16]. Digestive cells, also known as absorptive or secretion-reabsorption cells, have a

columnar shape and are the most abundant in the digestive gland of molluscs. They

present an intense endocytic activity, having many heterolysosomes where the

intracellular digestion of food particles occurs. They present lipid droplets and glycogen

granules in the cytoplasm, which can also be found in smaller amounts in basophilic cells

(Figure 3) [13, 14]. Basophilic cells, also known as secretory cells, calcium cells or crypt

cells, typically present a pyramidal shape and contain large amounts of rough

endoplasmic reticulum, a well-developed Golgi apparatus and secretory vesicles, being

considered responsible for secretion of enzymes involved in extracellular digestion

(Figure 3). They are responsible for the metal accumulation related with detoxification

[12, 14], but can also accumulate calcium in concentric structures known as calcium

concretions or spherites [17]. Excretory cells are considered a final stage of digestive

cells, exhibiting a large central vacuole and basal nucleus, and are responsible for the


15

excretion of residues into the tubule lumen. Thin columnar undifferentiated cells were

also reported in some species, which may give rise to digestive or basophilic cells [14].

Figure 3 - Schematic representation of the digestive gland cells in gastropods. mv – microvilli, ev – endocytic vesicle, ly –

lysosome, li – lipid droplets, nu – nucleus, sv – secretory vesicles, cc – calcium concretions (provided by A. Lobo-da-

Cunha, University of Porto).

Digestive gland cells are rich in peroxisomes. However, they present some

variability when both basophilic and digestive cells are compared. In basophilic cells,

these organelles are abundant in the middle and basal zones and present a nucleoid

with a variable shape between species. In digestive cells, peroxisomes are also found in

great number in the cytoplasm around vacuoles, typically with smaller size.

Morphological differences between digestive and basophilic cells peroxisomes were

reported in some gastropod species, thus suggesting functional differences [12, 13, 18].

Additionally, it was demonstrated that chemical pollutants can lead to morphological and

biochemical changes in peroxisomes, therefore being considered a valuable bioindicator

of pollution [19].

Despite much being known about the digestive system of gastropods, still many

doubts remain to be clarified. For example, how these cells and systems are regulated

accordingly to the animal’s activity and metabolic demands, is still poorly understood.

Another issue, is the fact that most field collected species are carried and maintained in

laboratory captivity in assumingly good conditions. However, some species of

gastropods are very sensitive to environmental conditions that can lead to drastic

changes of their metabolism, namely the digestive system [14]. Therefore, further

research is required for our better understanding of the metabolic regulation in these

organisms.


16

1.3. Oxidation of polyols and aromatic alcohols in gastropods

Ultrastructural analysis of the digestive gland of the sea slug Aplysia depilans

revealed many tubular structures inside the rough endoplasmic reticulum cisternae of

basophilic cells (Figure 4) [12]. Similar structures were also described in terrestrial

gastropods, such as the snail Helix pomatia [20] and the slug Arion empiricorum [21],

and in the digestive cells of the marine species Alderia modesta [22]. These structures

were called “mannosomes” since the enzyme mannitol oxidase, which catalyses the

oxidation of D-mannitol to produce the sugar mannose, releasing hydrogen peroxide

[23], is present in these structures, as shown by histochemical and biochemical

localization [24], and by purification methods [23, 25-28]. These studies confirmed the

capacity to metabolize D-mannitol in terrestrial [24-26] and marine [12] species.

Mannitol is a polyol present in plants, algae and fungi. Therefore, it was suggested

that herbivorous gastropods would get nutritional benefits from the ingestion of polyols,

present in plants [29] and algae [30] of their diet, due to the enzymes that turn these

compounds into sugars [24].

Figure 4 - [A] Ultrastructure of the basophilic cells of the digestive of A. depilans, showing tubular structures in the interior

of rough endoplasmic reticulum cisternae (*) [12]. [B] Frozen section of the digestive gland of A. depilans showing stained

basophilic cells (arrows) after the detection of mannitol oxidase activity [12].

More recently, the presence of mannitol oxidase in the digestive gland of several

herbivorous gastropods (Table S1), including terrestrial, marine and freshwater species,

was confirmed by activity assays [31]. In Patellogastropoda, Neritimorpha and

Vetigastropoda the activity of mannitol oxidase was not detected. However, mannitol

oxidase activity was detected in all herbivorous species belonging to Caenogastropoda


17

and Heterobranchia. The tubular structures inside rough endoplasmic reticulum

cisternae, typically associated with mannitol oxidase activity [12, 26], were found in

Aplysia depilans, while in Bulla striata, Cornu aspersum and Lehmannia valentiana, they

were found in smooth membrane cisternae. It was also reported that some of the species

expressing mannitol oxidase activity, such as Siphonaria pectinata and Onchidella

celtica, do not exhibit the typical tubular structures [31]. So, in what concerns marine

gastropods, mannitol oxidase is not always associated with intracisternal tubules even

in species with a high activity, such as O. celtica [31].

Another polyol with both nutritional and osmoregulation effects is the mannitol

isomer, sorbitol [32]. In previous studies (Table S1), sorbitol oxidation was only detected

in herbivorous species. In Caenogastropoda only Littorina littorea does not express

sorbitol oxidase activity. In Heterobranchia this activity is restricted to the Eupulmonata

clade. This data supported the hypothesis that mannitol oxidase can also use sorbitol as

substrate (data not published).

Another interesting finding about alcohol oxidases in the digestive gland of

terrestrial gastropods is their capacity to use aromatic alcohols as substrate [33, 34]. This

type of activity was first reported in the culture fluid of the lignolytic white-rot fungus

Polystictus versicolor [35]. Later, several groups have reported enzymes with the same

activity in a variety of other lignolytic fungi [36, 37]. Fungal aromatic alcohol oxidase

activity correlates with lignin degradation. In gastropods, aromatic alcohol oxidase show

some similarities in terms of pH optimum and substrate preferences [34]. Moreover,

herbivorous gastropods species can consume plant matter that has undergone some

lignification, or contains lignin precursors. Thus, it is likely that these species also have

this oxidase activity. Studies on aromatic alcohol oxidase in gastropods showed that this

enzyme has the highest activity towards cinnamyl alcohol, among several substrates

tested [34]. Surprisingly, our data (Table S1) show that all analysed gastropod species,

except O. celtica, present oxidase activity for cinnamyl alcohol (data not published).

In addition to polyols and aromatic alcohols, low molecular weight alcohols were

also tested. In some species, oxidation of ethanol was detected, however none of them

showed oxidation of methanol. Researchers frequently discuss whether the natural

intake of ethanol, in significant concentrations, originated an adaptation in the organisms

that can metabolize it, or if its capacity to oxidize ethanol is just a non-specific and

incidental response [38-40]. Studies using gastropods revealed that this activity is

present in Patella vulgata, in the carnivore caenogastropod Charonia lampas, and in

Heterobranchia, in the freshwater species P. corneus and in all terrestrial species, the

Stylommatophora (Table S1). However, with this data only it was not possible do

conclude if this capacity is an adaptation or an incidental feature (data not published).


18

1.4. Alcohol dehydrogenases

Oxidoreductases catalyse oxidation and reduction reactions where an electron is

transferred from a molecule to another. They can be divided into six categories:

oxygenases, reductases, peroxidases, oxidases, hydrolases and dehydrogenases. This

large group of oxidoreductases shows their importance and functional diversity in the

different organisms [41].

The roles of this group of enzymes are quite wide and comprises many

intermediary metabolic functions such as utilization and detoxification of ethanol and

xenobiotics in general, regulation of hormones and signalling molecules or just sensing

the redox status of the organism, thereby regulating vital cellular processes.

The oxidation of a substrate via dehydrogenase is characterized by the

transference of a hydrogen to an electron acceptor, such as NAD+ or FAD, in which the

substrate is losing electrons [42].

Alcohol dehydrogenases (ADH) are widely distributed among organisms of all

phyla and are responsible for the oxidation of alcohols into aldehydes or ketones [43].

There are three non-homologous NAD(P)+ -dependent alcohol dehydrogenase families

in animals. All of them evolved independently and have different structures and reaction

mechanisms [44, 45].

1.4.1. MDR - Type I ADHs

Type I ADH or “medium-chain” dehydrogenase/reductase (MDR) is the most

studied ADH superfamily in animals. Its functions include the metabolism of ethanol,

hydroxysteroids, lipid peroxidation products, formaldehyde, ω-hydroxy fatty acids and

norepinephrine, nitric oxide homeostasis, and retinoid transformation These are NAD+

and zinc-dependent enzymes, with one catalytic and one structural zinc atom, which

have between 350 and 375 amino acid residues. The region around and after the

structural zinc atom is the most divergent region in all ADH proteins [38, 46].

Based on sequence homology, catalytic activity and gene expression, these

enzymes have been sorted in eight different classes (I-VIII) in animals. In invertebrates,

there are only class III ADHs, in fish there are class I and III, in amphibian there are nine

or ten isozymes distributed by four or six classes, and in mammals there are six different

classes, but primates lack class VI ADH [38]. This multiplicity of ADH’s suggests that,

beyond their metabolic function and organism regulation, these enzymes play a role in


19

cellular defence against toxic alcohols and aldehydes, both endogenous and exogenous.

They appear to work together with cytochrome P450 and other oxidative systems, acting

without generation of potentially dangerous reactive oxygen species [47].

Class I Zn-dependent alcohol dehydrogenases

This class of enzymes is the most studied regarding the theme of ethanol-

metabolizing enzymes in different organisms, mainly due to its relevance for the alcoholic

tolerance [48]. They were originated by tandem duplication of class III ADH, 450 million

years ago, back in the origin of bony fish [49]. Since then, multiple independent gene

duplication of these enzymes has occurred in the different animal groups. The more

antique groups of vertebrates, fish and amphibians, have 3-5 and 3-6 class I ADH genes,

respectively. In mammals like cow, dog, mouse, and rat there is only on gene, but in

humans and other primates, there are three gene copies [50]. Additionally, a study to

trace overall patters within mammalian ADH genome, which was performed using data

from 15 genomes, revealed that this class of ADH is divided into three isoforms, ADH I

A, ADH I B and ADH I C [51].

In fish and amphibians, the appearance of these ADH isozymes is not related with

ethanol exposure, since they do not have any type of fruit in their diet. So these animals

have what is called a class I ADH ethanol availability-independent gene multiplicity [38].

In mammals, however, there is a broad array of studies on gene expression induced by

ethanol enriched diets [52], influence of alleles on ethanol-metabolizing processes and

its relation with the tolerance of the organism [48] and even how social-economic factors,

such as alcoholic beverages consumption [53] or pre/pro-industrial societies [54], may

be related with these alcohol-metabolizing enzymes occurrences [55].

Despite being highly known for its ethanol-metabolizing action, the list of substrates

for class I ADH can be quite extensive, from short to long aliphatic alcohols, retinol,

hydroxysteroids, bile acids (ADH I C), and hydroxytryptophol (serotonin metabolism). In

mammals, these enzymes are inhibited by pyrazole which has been effectively used to

distinguish class I ADH activity from other activities [51].

Class II Zn-dependent alcohol dehydrogenases

Traced in amphibians 360 million years ago [49], class II ADH is strongly related

with natural exposure of ethanol. Interestingly, in rodents it shows low affinity for ethanol

when compared with other alcohol dehydrogenases [56], but much higher catalytic


20

activity with retinol. This is a good evidence of the non-specific behaviour of this group

of enzymes, regarding its substrates [38].

In human, this class of ADH was isolated as pyrazole insensitive enzyme and is

active towards primary alcohols. In the oxidative pathway, the enzyme has been shown

to be the best hepatic retinol dehydrogenase [57]. In the reductive pathway class II ADH

has low activity for benzoquinones, is active towards lipid peroxidation products and has

a suggested redox-specific function in norepinephrine metabolism. However, the main

function for class II ADH is still an open question [51]

Class III Zn-dependent alcohol dehydrogenases

This is the only class of ADHs present in all eukaryotic organisms, and some

bacteria, being the most conserved ADH from all classes. It is considered the ancestral

form of zinc-containing ADHs from which the other ADHs have evolved. It has been

found in all tissues of animals who have their genome completely sequenced [43].

Compared to the other classes, ADH III has two additional activities. It is also

known as glutathione-dependent formaldehyde dehydrogenase and S-

nitrosoglutathione reductase, being expressed in all species containing glutathione [58,

59]. Regarding mammals, this ADH is the only one that has been traced in the central

nervous system [60, 61].

ADH III has very low affinity for ethanol as substrate [62], being suggested to be

involved in the first-pass metabolism of ethanol [63]. Since ethanol is present at low

concentrations in human diet under natural conditions, mainly as ingested ripe fruits, the

role of class III ADH in ethanol metabolism can be considered as incidental and not

adaptive from an evolutionary perspective [38]. However, class III ADH has high activity

for different ω-hydroxyfatty acid and is considered a formaldehyde scavenger in the

pathway where S-hydroxymethyl glutathione is the actual substrate of this enzyme [64].

Class IV Zn-dependent alcohol dehydrogenases

Described as gastric ADH and non-hepatic ADH [65], the origin of class IV ADHs

can be traced to marsupials, 176 million years ago [49]. This is another class of ADHs

that originated before the dietary ethanol being available by means of fruits fermentation

and, therefore, is non-ethanol metabolizing enzyme class [38].

Regarding the human gastric mucosa, this class of enzymes is responsible for the

maintenance of retinoic acid supply [66] and its malfunction is correlated with progression

of inflammation, atrophy and even intestinal metaplasia [67].


21

In several species, class IV ADH has not been identified since the region where

the gene encoding for this enzyme is located is poorly analysed in numerous genomes.

On other hand, in opossum for example, there are several isoforms of these enzymes

[51].

Class V and VI Zn-dependent alcohol dehydrogenases

Class VI ADHs also have origin in marsupial, 176 million years ago, and originated

class V ADHs, 76 million year later, by means of tandem duplication. Class V ADHs

appear to be restricted to placental mammals [49]. These functional equivalent enzymes

are known only in rat, which express two isoforms of ADH VI (A and B), and cow.

Primates lack class VI and mouse and dog lack class V ADHs, being the last one a

possible pseudogene in mouse [51]. However, neither human class V nor rat class VI

have been isolated as active enzymes, which is why their physiological role is still

unknown [68, 69].

Overall, these two classes of ADHs are exceptions and have been identified only

at DNA and mRNA levels. At the transcriptional level, different sizes of mRNA have been

identified for class V ADH, but no stable protein has been isolated [70].

Birds Class VII and amphibians class VIII Zn-dependent alcohol dehydrogenases

Class VII ADH has been found only in chicken with higher activity towards all-trans-

retinol and epiandrosterone. Amphibians class VIII ADH displays a strong preference for

NADP+ instead of NAD+ and higher activity with all-trans-retinaldehyde. Thus, it seems

that birds class VII and amphibians class VIII ADH do not play a role in the ethanol

metabolism [38].

1.4.2. SDR - Type II ADHs

Most short-chain dehydrogenases/reductases (SDH) have less than 250 amino

acid residues. However, one family, in a total of five, presents an extension until 350

residues. It was shown that about ¼ of the total described dehydrogenases are SDR.

These are present in insects, such as the genus Drosophila, but about ¾ of the total

SDR’s are of bacterial origin [42].


22

The amino acid sequence of the fruit fly Drosophila melanogaster ADH was

obtained, 36 years ago [71], and is clearly different from the amino acid sequences of

mammalians and yeast ADHs [72, 73]. At that time, several authors proposed the

existence of two different ADHs superfamilies [74], one being the short-chain ADHs from

insects, and the others long-chain ADHs, nowadays called medium-chain ADHs.

The type II ADHs have been extensively studied in the genus Drosophila and do

not share any structural relationship with zinc-dependent ADHs, except for the

coenzyme-binding domain, also known as Rossmann-fold [42]. All-common type I class

III ADHs are also expressed in this genus, in addition to type II. This last one is

responsible for most ethanol dehydrogenase activity in Drosophila [75], being considered

the unique fly enzyme involved in ethanol metabolism [76].

The analysis of ADHs in different families of flies, showed that the best substrates

for Drosophila ADH are secondary alcohols instead the primary ones [77]. Contrarily,

analysis of ADH1 and ADH2 from Ceratitis capitata and Anastrepha fraterculus showed

that ethanol and primary alcohols are better substrates than secondary alcohols [78, 79].

This again confirms the variability that these families of enzymes possess towards their

substrates. The origin and diversification of flies with different short-chain ADH genes

occurred shortly after the origin of angiosperms with fleshy fruits from the Lower

Cretaceous [38],which took place about 130-140 million years ago [80].

In vertebrates, the enzyme that shows more similarity to the type II ADH is the 15-

hydroxyprostaglandin dehydrogenase (PGDH), which also shows a broad phyletic

distribution in all animal groups, from the simplest ones, such as Cnidaria, to the most

complex, like the vertebrates. So, it can be assumed that short-chain ADH from insects

are derived by duplication from PGDHs, since these are of older origin [38].

1.4.3. LDR - Type III ADHs

Long-chain dehydrogenases/reductases (LDH) have more than 375 amino acid

residues and are iron dependent molecules that were initially identified in

microorganisms as ethanol oxidizing enzymes [38]. Later, new members of this

superfamily were found displaying different activities, such as: lactaldehyde reductase

and glycerol dehydrogenase in Escherichia coli [81, 82], butanol dehydrogenase in

Clostridium acetobutylicum [83], ethanolamine utilization protein in Salmonella

typhimurium [84], and 1,3-propanediol dehydrogenase in Klebsiella pneumoniae [85].


23

These enzymes conserve both an NAD(P)+ binding site and an iron-binding motif.

This last one has only two of the three conserved histidine residues, which make it unable

to determine the specific iron requirement for each enzyme [86].

In rats and humans, the gene encoding for the LDR ADHFe1 is expressed in adult

liver, kidney, thymus, spleen, hypothalamus and adipocytes [87]. ADHFe1 displays

activity as hydroxyacid-oxoacid transhydrogenase (HOT), which catalyses the

conversion of -hydroxybutyrate into succinic semialdehyde with subsequent reduction

of α-cetoglutarate [86]. -Hydroxybutyrate is an energy regulator that promotes the

release of growth hormone [88], and was used illegally by athletes as a performance-

enhancing drug [89]. This data identifies ADHFe1 as an athletic-performance gene that

was positively selected during 400 years in Thoroughbred horses [90].

1.4.4. Polyol Dehydrogenases

Polyols or alditols are acyclic polyfunctional alcohols, also known as sugar

alcohols, that derive from aldoses or ketoses by reduction of the carbonyl group. They

are specially abundant in algae, fungi and plants [91]. Because of their physico-chemical

properties, these alcohols can be seen as simple carbohydrates. In animals, polyols are

important reserves of metabolites that regulate the density and viscosity of some

biological fluids such as blood, lymph and intercellular fluid [92]. These metabolic by-

products have also been associated with an increase of osmotic pressure that provides

a decrease of fluidity of plasma membranes by glucose derivatives, increasing the

cellular resistance to oxidative stress and some toxic agents [93]. The osmotic regulation

by these compounds was also reported in plants and algae [29, 30].

With a broad array of applications, polyols and their derivatives play an important

role in the production of everyday-products. In health industry, compounds like sorbitol

or xylitol are used as sugar substitutes in patient’s nutrition, allowing a better control of

problems regarding oral health and even diabetes control. Also, some polyol esters are

used as emulsifying agents, explosives and pharmaceuticals. Therefore, numerous

dehydrogenases with different substrates specificities have been characterized along the

years, like those involved in the interconversion of D-mannitol/D-fructose or the in vivo

oxidation of sorbitol to fructose by sorbitol dehydrogenase (SDH) [91].

Described as an additional pathway in glucose metabolism [94], the polyol/sorbitol

pathway (Figure 5) involves aldose reduction (ALR2) that convert aldoses into their

corresponding ketoses, and then SDH [95]. This pathway functions as a bypass to


24

glycolysis and the pentose phosphate pathway that was used as therapeutic tool in the

control of hyperglycaemia of some diseases, such as diabetes [96, 97].

Figure 5 - Scheme of the polyol pathway of the glucose metabolism. Conversion of glucose to sorbitol by aldose reductase,

and sorbitol to fructose by sorbitol dehydrogenase [98].

Sorbitol dehydrogenase (Figure 6), or L-iditol-2-dehydrogenase, is a 155 kDa

tetrameric protein, with each subunit having around 38 kDa. It has a total of 356 amino

acid residues and a single zinc atom, therefore belonging to the type I medium-chain,

zinc-dependent dehydrogenase/reductase family, but not fitting in any of the eight ADH

classes early described in animals [91]. It is homologous to the alcohol dehydrogenase,

which has a conserved zinc ligand motif that contains two cysteine residues, one

histidine residue and a hydrophobic substrate-binding pocket. The main difference

between ADH and SDH is the replacement of the third Zn ligand in ADH, a cysteine, by

a glutamine residue in SDH [98].

Figure 6 - Structure of sheep sorbitol dehydrogenase (http://www.rcsb.org/pdb/explore/explore.do?structureId=3QE3).

ALR2 SDH

Glucose Sorbitol Fructose

NADPH NAD+ NADP+ NADH


25

Sorbitol dehydrogenase is the second enzyme of the polyol pathway (Figure 5) and

catalyses de conversion of sorbitol to fructose. In the first step of this pathway, glucose

is converted to sorbitol by ALR2, a NADPH-dependent aldose reductase. SDH is widely

distributed in almost all mammalian tissues [99-101]. The coenzyme NADH was found

to bind more tightly to SDH than NAD+, at both high and low pH values [102].

The increased flux of glucose through the polyol pathway, by the action of ALR2,

in a hyperglycaemic state, decreases the NADPH/NADP+ ratio. Thereby, the reduction

of glutathione disulphide is impaired leading an oxidative stress. Additionally, the

increase of sorbitol flux through this pathway, because of the increased SDH activity,

may decrease the cytosolic NAD+/NADH ratio resulting in the block of the glycolytic

pathway at the level of glyceraldeyde-3-phosphate dehydrogenase (GAPDH). The

resulting imbalances of these redox states will further affect several other metabolic

pathways [98].

Several studies highlight the importance of these enzymes in the organism

homeostasis, or even as tool for diseases treatment. Nevertheless, other biochemical

imbalances can lead to its overstimulation. Such is the case of the deleterious effect of

sorbitol accumulation on various tissues, because of an overstimulated ALR2 activity

[103]. Stimulators of this pathway include haloalcohols and detergents that will alter the

catalytic site, resulting in a more flexible configuration, aiding the release of NADH from

the coenzyme binding site, and therefore originating a faster rate-limiting dissociation

process of SDH, when in the presence of high concentrations of sorbitol [104].

Similar maximum velocities with various polyol substrates have been previously

reported for human [105] and rat [106] liver SDH, and human [101], rat and bovine [107]

brain SDH. For many years, there was a lack of consensus in the literature regarding de

substrate specificity of SDH. In a first approach, it was suggested that polyol substrates

of SDH have the D-cis-2,4-dihydroxy (2S,4R) configuration, such is the case of sorbitol

and a primary hydroxyl group adjacent to the oxidation site at C2 [91]. Other studies [91]

have shown that SDH catalyses de reversible NAD-linked oxidation of most polyols,

including D-mannitol and galactitol, the 2- and 4-epimers of sorbitol, which do not

possess that specific configuration, into their corresponding ketones. Therefore, the

supposed D-cis-2,4-dihydroxy requirement for the polyol substrates of SDH is not

absolute [108]. Table 1 shows a list of possible substrates for human liver SDH.


26

Table 1 - Compatible substrates for human liver sorbitol dehydrogenase [105]

substrate relative rate

sorbitol 100

glycerol 6

erythritol 4

L-threitol 110

ribitol 85

xylitol 103

L-arabinitol 29

D-mannitol 8

galactitol 11

Some reports show that mammalian liver SDH can also use aromatic secondary

alcohols as substrate [109]. Overall, SDH does not bind substrates with C2 configuration

of a polyol and the length of the carbon chain determines the degree of enzymatic

discrimination. The C2 hydroxyl group is coordinated to the Zn atom while the primary

hydroxyl group and the substituent sits in a polar region and in a hydrophobic area,

respectively [91].

1.5. Alcohol and polyol dehydrogenases in molluscs

Molluscs are the second major phylum of multicellular animals. However, that fact

is not reflected in the number of studies of alcohol and polyol dehydrogenases in these

organisms.

A study with cephalopods molluscs (octopus, squid and cuttlefish) showed the

presence of only class III alcohol dehydrogenase, with a wide tissue distribution. This

enzyme behaves like the other class III alcohol dehydrogenases identified in other

organisms, with very low affinity for ethanol and high glutathione-dependent

formaldehyde dehydrogenase activity, proving that multicellular organisms can survive

without ethanol active alcohol dehydrogenases. No other class of alcohol

dehydrogenase enzyme was found in the cephalopod species or organ tissues tested.

In fact, cephalopods were the first multicellular organisms known without significant

amounts of alcohol dehydrogenase forms capable of metabolizing low molecular weight

alcohols, such as ethanol [110].

Biomphalaria glabrata is a gastropod that has been extensively studied in the

matter of public health. To develop mechanisms for the suppression of natural


27

populations of this species, several strategies of genetic manipulation have been used.

Some techniques only produce effect after several generations, so it is important to

understand the polymorphisms and inheritance pattern of certain enzymes. Therefore,

there are some studies regarding alcohol dehydrogenases in these species [111].

Additionally, due to the fact of being a subject of extensive investigation, this freshwater

snail is one of the few molluscs with its genome fully sequenced.

Class III alcohol dehydrogenase of the marine species Nucella lapillus was studied

in the context of environmental contamination with endocrine disrupting chemicals.

Chemicals of high persistence and bioaccumulation are used in agriculture, industry and

antifouling paints for boats and fishing nets, and can affect the endocrine system of

several organisms, leading to deregulation of important physiological functions. Studies

with ascidians showed that class III ADH gene expression was strongly downregulated

upon tributyltin exposure. The disruptive consequences of this pollutant in

caenogastropods, such is the case of N. lapillus, is the imposex phenomenon by altering

retinoid X receptors activation. That leads to the development of male secondary sexual

organs in females. Therefore, these animals play a role of major importance as

bioindicators of water contamination. However, in this study, although tributyltin induced

imposex after two-month exposure, class III ADH expression of N. lapillus was not

significantly altered. On the other hand, retinol significantly downregulated class III ADH

levels in female gonads [112].

Regarding organism homeostasis, sorbitol dehydrogenase plays a key role in the

regulation of sorbitol concentration and therefore the viscosity of biological fluids in the

freshwater snail Viviparus viviparus. This way, the organism can adjust some physical

parameters of its liquid phase, such as decreasing the freezing temperature. This stress-

induced cryoprotecting feature is particularly useful for poikilothermic animals allowing

them to resist to condition of low temperature [32],

1.6. Aims of this work

At this point it is quite evident the lack of knowledge about the alcoholic

dehydrogenases in molluscs. Therefore, the objective of this work was to study how

aromatic alcohol and polyol dehydrogenases are distributed among the different clades

of gastropods and how that can be related with their dietary habits. This work also aimed

to isolate and identify the enzyme(s) responsible for the metabolization of specific

substrates in the true limpet Patella vulgata.


28

2. Materials and Methods

2.1. Species and sampling sites

Most of the marine species used for this study were collected in the North coastal

region of Portugal: Patella vulgata, Phorcus lineatus (=Monodonta lineata), Gibbula

umbilicalis, Calliostoma zizyphinum, Nassarius reticulatus, Charonia lampas, Nucella

lapillus, Aplysia depilans, Ocenebra erinaceus and Siphonaria pectinata. From Ria de

Aveiro, a coastal lagoon connected to the Atlantic Ocean on the central coastline of

Portugal, was obtained the common periwinkle Littorina littorea. The sea slugs Aglaja

tricolorata, Pleurobranchaea meckelii and Armina maculata were collected in Troia

beach near Setubal. The marine species Hexaplex trunculus, Doriopsilla areolata, Bulla

striata, Haminoea orbignyana and Onchidella celtica were obtained on the South coast

of Portugal (Algarve). The freshwater snails Neritina natalensis, Pomacea bridgesii, and

Marisa cornuarietis were purchased in aquarium shops. Planorbarius corneus

exemplars, also a freshwater snail, were offered by Centro de Educação Ambiental das

Ribeiras de Gaia (CEAR). The terrestrial species Cornu aspersum (=Helix aspersa),

Lehmannia valentiana and Arion ater were collected in the North of Portugal. The

taxonomic position, habitat and diet of these species are included in Table 2.

2.2. Preparation of protein extracts

The digestive gland of each animal was removed and homogenized at 1000 rpm

in a Potter-Elvehjem homogenizer at 4 ºC, using an homogenizing medium for marine

invertebrates (50 mM Tris-HCl buffer pH 7.4, 500 mM sucrose, 150 mM KCl, 1 mM EDTA

and 1 mM PMSF), in an average concentration 0,06 g/ml of digestive gland (adapted

from [113, 114]). The homogenates were then sonicated in three cycles, 15 s each, in

ice-water bath (Bandelin – Sonorex RK 100H sonicator) and centrifuged at 1000 g for 5

min. at 4 ºC. The supernatants were used to determine enzyme activities.


29

Table 2 - Taxonomy, habitat and diet of the 26 species of gastropods included in this study. Ma - marine, Fw - freshwater, Te - terrestrial

Gastropod clades Superfamily Family Species Habitat / Diet

Patellogastropoda Patelloidea Patellidae Patella vulgata Ma / Herbivore

Neritimorpha Neritoidea Neritidae Neritina natalensis Fw / Herbivore

Vetigastropoda Trochoidea Trochidae

Phorcus lineatus Ma / Herbivore

Gibbula umbilicalis Ma / Herbivore

Calliostomatidae Calliostoma zizyphinum Ma / Carnivore

Caenogastropoda

Hypsogastropoda

Buccinoidea Nassariidae Nassarius reticulatus Ma / Carnivore

Tonnoidea Ranellidae Charonia lampas Ma / Carnivore

Muricoidea Muricidae

Nucella lapillus Ma / Carnivore

Hexaplex trunculus Ma / Carnivore

Ocenebra erinaceus Ma / Carnivore

Littorinoidea Littorinidae Littorina littorea Ma / Herbivore

Ampullarioidea Ampullariidae Pomacea bridgesii Fw / Herbivore

Marisa cornuarietis Fw / Herbivore

Heterobranchia

Nudipleura

Pleurobranchomorpha Pleurobranchoidea Pleurobranchidae Pleurobranchaea meckelii Ma / Carnivore

Nudibranchia Doridacea Phyllidioidea Dendrodorididae Doriopsilla areolata Ma / Carnivore

Cladobranchia Arminoidea Arminidae Armina maculata Ma / Carnivore

Euopisthobranchia Cephalaspidea

Philinoidea Aglajidae Aglaja tricolorata Ma / Carnivore

Haminoeoidea Haminoeidae Haminoea orbignyana Ma / Herbivore

Bulloidea Bullidae Bulla striata Ma / Herbivore

Anaspidea Aplysioidea Aplysiidea Aplysia depilans Ma / Herbivore

Pan

pu

lmo

nat

a Siphonarioidea Siphonaridae Siphonaria pectinata Ma / Herbivore

Hygrophila Planorboidea Planorbidae Planorbarius corneus Fw / Herbivore

Eupulmonata

Systellommatophora Onchidioidea Onchidiidae Onchidella celtica Ma / Herbivore

Stylommatophora

Helicoidea Helicidae Cornu aspersum Te / Herbivore

Arionoidea Arionidae Arion ater Te / Omnivore

Limacoidea Limacidae Lehmannia valentiana Te / Herbivore

http://en.wikipedia.org/wiki/Nassariidae

http://en.wikipedia.org/wiki/Muricidae

http://en.wikipedia.org/wiki/Ampullariidae

http://en.wikipedia.org/wiki/Hygrophila_(Gastropoda)


30

For preparation of cytosolic fractions of the digestive gland of Patella vulgata, total

homogenates with 10% digestive gland tissue (4 g of digestive gland in a 40-ml

homogenate), were obtained using a salt-free homogenizing medium (50 mM Tris-HCl

buffer pH 7.4, 500 mM sucrose, 1 mM EDTA and 1 mM PMSF). Samples were

centrifuged at 15 000 g for 15 min. at 4 ºC and then at 100 000 g for 1 h, at 4 ºC (Thermo

Scientific; Sorvall Lynx 6000, T29 - 8x50 ml rotor).

The Bradford method with bovine serum albumin standards was used for protein

quantification [115].

2.3. Enzyme activities

The specific activity of dehydrogenases was determined by using

spectrophotometric assays.

Twenty µl of cytosolic sample were added to 980 µl incubation medium (50 mM

glycine-NaOH buffer pH 10, 5 mM NAD+) supplemented with 50 mM D-sorbitol, 50 mM

D-mannitol or 25 mM cinnamyl alcohol (adapted from [110]). The formation of NADH

(ɛ340 nm = 6.22 mM-1 cm-1) [29] was followed by measuring O.D.340 at 25 ºC, during 200 to

300 sec. A reference cuvette with the incubation medium without sample was used. The

presented values of activity in the total homogenates are mean of at least 5 independent

experiments, and were calculated by using the following equation: Activity (nmol-1ml-

1min-1) = (∆ O.D.340 min. / 0.00622 µM-1) × 50. These assays were done using a double-

beam spectrophotometer (Jenway 68000 UV/Vis.), coupled to a water recirculating

system for temperature regulation in the cuvette.

The activity of specific dehydrogenases was also studied in situ, after separation

of the cytosolic fraction proteins (60-100 µg) by native gel electrophoresis, using 7.5%

non-denaturant polyacrylamide gels. After electrophoresis, the gels were the incubated

in a medium containing 50 mM Tris-HCl buffer pH 8.0, 0.01 mg/ml phenazine

methosulphate, 5 mM NAD+, 0.10 mg/ml NBT (nitro blue tetrazolium chloride) and the

alcoholic substrate (above mentioned concentrations), until the appearance of the

bands. In the control gel the same process was executed but without the alcoholic

substrate (adapted from [116]).


31

2.4. Affinity chromatography

The sample (one column volume; 5 ml) was loaded on a Blue Sepharose CL-6B

column and the flow through was recovered. The column was washed with five volumes

(25 ml) of 0.1 M Tris-HCl pH 7.4 buffer. The first 2 volumes were recovered for the control

of activity lost at this step. Then, proteins bound to the column were eluted with one

column volume of increasing concentrations of NAD+ solution (0.05 M, 0.1 M, 0.2 M).

Each of these fractions was recovered independently. Column regeneration was

performed by washing with 2 column volumes, 3 to 5 times, of alternating high (0.1 M

Tris-HCl pH 8.5, 0.5 M NaCl) and low (0.1 M Sodium Acetate pH 4.5, 0.5 M NaCl) buffers.

The NAD+ eluted fractions were concentrated in a Vivaspin 500 Protein Concentrator (6

ml, MWCO 10 000, GE Healthcare Life Sciences) to a final volume around 2.5 ml, due

to its high dilution.

2.5. Ion exchange chromatography

For cation exchange chromatography two columns were used: HiTrap SP HP, 1

ml, controlled by ÄKTAprime plus, a low pressure chromatographic system (GE

Healthcare Life Sciences) and Mono Q 4.6/100 PE controlled by BioLogic DuoFlow™, a

medium pressure chromatographic system (Bio-Rad).

Prior to sample injection, the columns were washed with volumes of alternating low

salt (20 mM Tris-HCl pH 7.4), high salt (20 mM Tris-HCl pH 7.4 1 M NaCl) and low salt

buffers. Concentrated 10 mM NAD+ fractions obtained by affinity chromatography were

diluted with 20 mM Tris-HCl pH 7.4 buffer, for conductivity adjustment, before being

injected in these systems with the aid of a peristaltic bomb. After injection, the column

was washed one more time with low salt buffer for non-bound protein removal. The

columns were then eluted with an ascending and continuous NaCl gradient from 0.05 M

to 1 M, at a flow rate of 1 ml per sec.

2.6. SDS-polyacrylamide gel electrophoresis

To study the protein profile of the samples resulting from the chromatographic

processes, SDS-polyacrylamide precast gels were used (Bio-Rad 12% Mini-


32

PROTEAN® TGX™ Precast Protein Gels). Two hundred µl of sample were mixed with

25 µl of 100% TCA, incubated on ice for 5 min., and centrifuged at 10 000 g for 5 min.

The pellet was washed abundantly with acetone to remove the acid, dissolved in 20 µl

of sample buffer (0.25 M Tris-HCl pH 6.8, 8% SDS, 40% glycerol, 20% 2-

mercaptoethanol) and heated at 90 ºC for 5 min. The samples and molecular weight

standards solution (GE Healthcare Life Sciences LMW-SDS Marker Kit: Phosphorylase

b, 97 kDa; Albumin, 66 kDa; Ovalbumin, 45 kDa; carbonic anhydrase, 30 kDa; trypsin

inhibitor, 20.1 kDa; -lactalbumin, 14.4 kDa) were then separated by electrophoresis and

the proteins were stained for 2 hours, using 0,025 M Coomassie Brilliant Blue R-250,

40% methanol, 7% acetic acid). The gel was destained using a 50% methanol, 10%

acetic acid solution.

2.7. Protein sequencing

Coomassie-blue stained protein bands were excised from a SDS-polyacrylamide

gel and analysed by mass spectrometry (MS). Proteins were in gel digested with trypsin

(Promega, USA) and the peptides were extracted using a 2.5% trifluoroacetic acid

solution. Protein digests were desalted and concentrated using C18 ZipTips (Millipore,

USA) and crystallized onto a MALDI plate using α-Cyano-4-hydroxycinnamic acid as a

matrix. Samples were analysed using the 4800 Plus MALDI-TOF/TOF (SCIEX, USA)

[117]. Protein identification was performed by Peptide Mass Fingerprint with Mascot

software (Matrix Science, UK). Some peptides were further characterized by MS/MS

sequencing. The identification of peptides from acquired MS+MS/MS spectra was

performed using a Patella vulgata database (available at:

https://ora.ox.ac.uk/objects/uuid:6471e7d4-dd34-4eb3-883f-a5c438f11741). A Blastp of

the identified proteins was performed at the NCBI database.


33

3. Results

3.1. Enzymatic activities

To analyse the distribution of polyol and aromatic alcohol dehydrogenases in five

major clades of gastropods, the activity of these enzymes was quantified using total

homogenates of the digestive gland of 26 species of gastropods.

Regarding the dehydrogenase for the polyols D-mannitol and D-sorbitol (Table 3),

almost all species presented activity for both substrates, with a few exceptions. The

species B. striata and P. meckelii only presented activity for sorbitol whereas the

carnivore vetigastropod C. zizyphinum, the caenogastropods N. reticulatus and H.

trunculus, and the Heterobranchia A. maculata, did not exhibit dehydrogenase activity

for the tested polyols.

Among the species with dehydrogenase activity for both polyols (Table 3), P.

vulgata presented high activity values for both substrates. However, sorbitol

dehydrogenase activity in M. cornuarietis was more than two-fold higher comparing to

that observed in P. vulgata. It was observed that only P. vulgata, P. lineatus and G.

umbilicalis did not present higher dehydrogenase activity for sorbitol than mannitol.

Overall, there was a tendency for higher sorbitol dehydrogenase activities than mannitol

dehydrogenase activities, and the carnivores presented lower dehydrogenase activities

for these substrates than the herbivores.

The dehydrogenase activity for cinnamyl alcohol, representing the aromatic

alcohols, was present in all species, without exceptions (Table 3), with the marine

herbivore Haminoea orbignyana from Heterobranchia exhibiting higher levels. However,

the carnivore N. lapillus also presented a high dehydrogenase activity for cinnamyl

alcohol. Overall, the four studied freshwater species presented low activity values.

Regarding dehydrogenase activities towards the low molecular weight alcohols,

ethanol dehydrogenase activity was only detected, at low levels, in two herbivorous

Eupulmonata species, the marine O. celtica and the terrestrial C. aspersum (Table 3).

No species presented methanol dehydrogenase activity.


34

Table 3 - Activity of dehydrogenase in the digestive gland of gastropods. Values are mean ± SD of at least 5 independent experiments (nmol.min-1. g-1 ± SD). Shaded lines correspond to carnivorous

species.

Species Mannitol Sorbitol Cinnamyl alcohol Ethanol

Patellogastropoda Patella vulgata 1,836.1 ± 368.6 1,644.3 ± 723.0 527.6 ± 154.8 -

Neritimorpha Neritina natalensis 870.2 ± 455.9 900.5 ± 356.4 215.4 ± 67.5 -

Vetigastropoda

Phorcus lineatus 1,214.4 ± 340.8 536.6 ± 211.2 1,205.8 ± 424.3 -

Gibbula umbilicalis 1,322.9 ± 244.5 818.9 ± 320.0 560.7 ± 125.2 -

Calliostoma zizyphinum - - 643.6 ± 197.7 -

Caenogastropoda

Nassarius reticulatus - - 1,385.0 ± 392.8 -

Charonia lampas 161.9 ± 60.4 418.4 ± 115.0 824.8 ± 240.5 -

Nucella lapillus 96.2 ± 97.1 381.4 ± 164.8 2,048.5 ± 670.5 -

Hexaplex trunculus - - 894.5 ± 208.9 -

Ocenebra erinaceus 109.1 ± 122.5 254.1 ± 332.5 1,007.4 ± 515.7 -

Littorina littorea 510.2 ± 157.5 944.9 ± 293.8 932.2 ± 432.7 -

Pomacea bridgesii 379.6 ± 65.3 741.0 ± 191.8 181.9 ± 54.5 -

Marisa cornuarietis 1,428.9 ± 232.1 4,095.5 ± 603.3 503.9 ± 75.2 -

Heterobranchia

Pleurobranchaea meckelii - 60.7 ± 68.0 498.0 ± 89.3 -

Doriopsilla areolata 202.0 ± 111.0 327.3 ± 141.9 117.2 ± 39.8 -

Armina maculata - - 1,109.5 ± 279.6 -

Aglaja tricolorata 133.6 ± 71.5 222.3 ± 89.4 1,482.8 ± 205.9 -

Haminoea orbignyana 224.0 ± 32.7 521.3 ± 139.5 2,643.5 ± 996.5 -

Bulla striata - 192.7 ± 82.6 2,017.4 ± 702.2 -

Aplysia depilans 131.5 ± 53.5 530.2 ± 150.1 727.2 ± 248.7 -

Siphonaria pectinata 333.8 ± 116.3 460.2 ± 79.7 1,013.7 ± 372.0 -

Planorbarius corneus 281.3 ± 140.7 847.9 ± 288.6 209.2 ± 60.6 -

Onchidella celtica 623.8 ± 235.5 865.7 ± 157.4 1,604.7 ± 389.0 307.9 ± 186.9

Cornu aspersum 546.2 ± 207.9 1,171.1 ± 335.8 1,329.6 ± 282.5 300.8 ± 137.2

Arion ater 651.6 ± 309.6 1,058.7 ± 501.1 1,264.5 ± 417.6 -

Lehmannia valentiana 437.4 ± 103.3 1,408.9 ± 319.2 978.1 ± 231.0 -


35

The activity of dehydrogenases for the polyols D-mannitol and D-sorbitol and for

the aromatic cinnamyl alcohol was also assessed in situ, after native gel electrophoresis

(Figure 7), using four marine herbivorous species: P. vulgata, P. lineatus, L. littorea and

S. pectinata. Regarding the polyols dehydrogenases, a similar overall profile was

observed in all species. For P. vulgata, L. littorea and S. pectinata, one main band of

enzyme activity was detected. Interestingly, sorbitol dehydrogenase activity was higher

when comparing with mannitol dehydrogenase. In L. littorea a second well defined band

appeared for each polyol, again with higher intensity for sorbitol. In P. lineatus, a main

band of sorbitol dehydrogenase activity was detected, whereas for mannitol

dehydrogenase two bands appeared, one of them with lower relative migration distance.

Figure 7 - Native electrophoresis gel sections with in situ detection of the activity of dehydrogenases of several alcoholic

substrates, in four marine herbivorous species.

Patella vulgata Phorcus lineatus

Littorina littorea Siphonaria pectinata

Mannitol Sorbitol Cinnamyl A. Mannitol Sorbitol Cinnamyl A.

Mannitol Sorbitol Cinnamyl A. Mannitol Sorbitol Cinnamyl A.


36

The profiles for cinnamyl alcohol dehydrogenase (Figure 7) revealed only one band

of enzyme activity in P. vulgata, while in the other species more bands were detected.

The faster migration band had a higher intensity, except for P. lineatus, in which the band

with lower relative migration distance exhibited a slightly higher intensity. L. littorea was

the species that presented more bands associated with cinnamyl alcohol dehydrogenase

activity (three).

3.2. Protein isolation and identification

Due to the significant amounts of alcoholic dehydrogenase activities, especially for

polyols, observed in previous studies (Table 3), the facility in obtaining good amount of

digestive gland per animal for sample preparation, and the fact of being an all-year

abundant species of easy collection, the species P. vulgata was used for the isolation

and identification of dehydrogenases.

First, proteins (cytosolic fraction) of this species were separated by affinity

chromatography, using Blue Sepharose CL-6B, which binds most enzymes requiring

adenyl-containing cofactors including NAD+. Proteins were eluted with NAD+ (5-20 mM)

and dehydrogenase activities were determined in the different fractions obtained (Table

4). The results show that the polyols dehydrogenases were more concentrated in the

washing steps after the sample being loaded in the column, and in the fractions eluted

with 5 and 10 mM NAD+. On the other hand, the enzymes responsible for cinnamyl

alcohol dehydrogenase were eluted with 20 mM NAD+. Thus, to obtain a fraction

enriched in polyol dehydrogenases for subsequent chromatographic tests, the Blue

Sepharose CL-6B column was eluted with 10 mM NAD+ (Table 5).

After Vivaspin concentration, the 10 mM NAD+ affinity chromatography fraction

was loaded in the HiTrap SP HP chromatography column and proteins were eluted using

NaCl gradient (Figure 8). The chromatogram shows 2 major protein peaks around the

fraction 22 and 25 (blue line). The activity for the alcoholic substrates was measured in

fractions 4 (flow through), 22, 25 and 27 (Table 6). Since high specific activities for

polyols dehydrogenase were identified in the fractions 22 and 25, samples 22 to 26 were

analysed by SDS-PAGE (Figure 9).


37

Table 4 - The cytosolic fraction of the digestive gland of Patella vulgata was loaded on a Blue Sepharose CL-6B column,

the flow-through was recovered and the column washed with buffer. Proteins were eluted with NAD+ solutions of

increasing concentrations (5 mM, 10 mM and 20 mM). The column was then washed with regeneration buffers.

Dehydrogenase activities for polyols (D-mannitol and D-sorbitol) and cinnamyl alcohol (aromatic) in the fractions were

assayed by spectrophotometry. A/ml – activity per ml of fraction; A/mg – activity per mg of protein (specific activity); A

total – activity of total fraction volume.

Mannitol Sorbitol Cinnamyl Alcohol

Fractions A/ml A/mg A total A/ml A/mg A total A/ml A/mg A total

Cytosol 152.89 28.93 764.45 103.46 19.57 517.30 47.27 8.94 236.35

Flow-Through 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00

Wash 5.43 5.43 135.75 1.29 1.29 32.25 0.77 0.77 19.25

NAD+ 5 mM 9.04 456.57 45.20 4.70 237.37 23.50 0.00 0.00 0.00

NAD+ 10 mM 15.92 151.19 79.60 25.20 239.32 126.00 3.14 29.82 15.70

NAD+ 20 mM 7.11 40.54 35.55 12.42 70.81 62.10 10.73 61.17 53.65

Wash pH 8,5 1.17 2.16 5.85 5.79 10.67 28.95 9.65 17.79 48.25

Wash pH 4,5 0.00 0.00 0.00 0.00 0.00 0.00 7.45 12.89 37.25

Table 5 - The cytosolic fraction of the digestive gland of Patella vulgata was loaded on a Blue Sepharose CL-6B column,

the flow-through was recovered and the column washed with buffer. Proteins were eluted with two NAD+ solutions of

different concentrations (10 mM and 20 mM). The column was then washed with regeneration buffers. Dehydrogenase

activities for polyols (D-mannitol and D-sorbitol) and cinnamyl alcohol (aromatic) in the fractions were assayed by

spectrophotometry. A/ml – activity per ml of fraction; A/mg – activity per mg of protein (specific activity); A total – activity

of total fraction volume.



Cytosol 207.88 39.03 1039.40 148.07 27.80 740.35 99.60 18.70 498.00

Wash 22.19 33.34 610.23 5.14 7.72 141.35 0.00 0.00 0.00

NAD+ 10 mM 136.49 457.25 341.23 102.01 341.74 255.03 27.01 90.49 67.53

NAD+ 20 mM 22.43 184.91 56.08 45.10 371.81 112.75 20.74 170.98 51.85

Wash pH’s 1.71 4.40 17.10 2.15 5.54 21.50 26.37 67.93 263.70

Table 6 - 10 mM NAD+ fraction from affinity chromatography loaded on a HiTrap SP HP column. Proteins eluted with an

ascending and continuous NaCl gradient (0.05 M to 1 M), at a flow rate of 1 ml per sec. (Figure 8). Distribution of

dehydrogenase activities for polyols (D-mannitol and D-sorbitol) and cinnamyl alcohol (aromatic) in some fractions were

assayed by spectrophotometry. A/ml – activity per ml of fraction; A/mg – activity per mg of protein (specific activity); A

total – activity of total fraction volume. Fractions: no.4 – flow-through representative, no. 22, 25 and 27 – protein peaks

representatives.



no. 4 2.19 260.71 2.19 0.70 83.33 0.70 0.00 0.00 0.00

no. 22 17.85 1593.75 17.85 2.89 258.04 2.89 0.00 0.00 0.00

no. 25 35.53 1015.14 35.53 49.03 1400.86 49.03 11.09 316.86 11.09

no. 27 4.10 259.49 4.10 4.82 305.06 4.82 0.00 0.00 0.00


38

Figure 8 - Ion exchange chromatogram of 10 mM NAD+ fraction from affinity chromatography with HiTrap SP HP column. Proteins eluted with an ascending and continuous NaCl gradient (0.05 M to 1

M), at a flow rate of 1 ml per sec. (blue) protein detection (abs 280 nm); (green) high salt solution concentration; (red) conductivity.


39

Figure 9 - SDS-polyacrylamide gel showing the enzymatic profile of some ion exchange chromatography fractions

obtained with HiTrap SP HP column (Figure 8). Proteins identified by MS/MS using a database of Patella vulgata: 1 –

glutathione-S-transferase mu class; 2 – no statistical significant identification; 3 – D-arabinitol dehydrogenase 1; 4 –

cytosolic malate dehydrogenase; 5 – sorbitol dehydrogenase-like protein (Table 7).

Some bands of interest were excised from the gel and proteins were analysed by

mass spectrometry (MS/MS) and identified recurring to a Patella vulgata database (Table

7). The following proteins were identified in fraction 22, where the specific activity for

mannitol was much higher than for sorbitol: glutathione-S-transferase and D-arabinitol

dehydrogenase. The analysis of another band around 40 kDa did not give a statistically

valid identification. In fraction 25, where the specific activity is high for both substrates

but higher for sorbitol (Table 6), a cytosolic malate dehydrogenase and a sorbitol

dehydrogenase-like protein were identified (Table 7). Also, the specific activity for

cinnamyl alcohol was higher in fraction 25.

1

2

3

4

5

14

20

30

45

66

97

LMW

kDa 22 23 24 25 26

Fractions


40

Table 7 - Proteins of interest (Figure 9) were analysed by mass spectrometry (MS/MS) and identified using a Patella vulgata database. Blastp of the identified proteins was performed at the NCBI

database. Mascot search result scores greater than 65 are significant (p<0.05).

Mascot Search NCBI Blastp

Band Accession MW

(kDa) Score

No. Peptides Identified

No. Amino Acids

Specific hits Organism Score Identity

(%)

1 CL19536Contig1 29.231 416 54 252

Hypothetical protein LOTGIDRAFT_233779 Lottia gigantea 357 77

Glutathione S-transferase mu class

Crassostrea ariakensis 322 69

Reishia clavigera 317 71

Cipangopaludina cathayensis 310 69

2 CL63Contig2 145.676 41 33

3 CL9544Contig1 44.49 665 49 399

Hypothetical protein LOTGIDRAFT_188066 Lottia gigantea

452 60

Hypothetical protein LOTGIDRAFT_159773 450 58

D-arabinitol dehydrogenase 1 Neodiprion lecontei 308 45

Ciona intestinalis 303 44

4 CL31882Contig1 37.963 1160 59 345


Cytosolic malate dehydrogenase

Lottia scabra 516 83

Lottia digitalis 509 81

Lottia scutum 508 81

Lottia austrodigitalis 506 81

Lottia pelta 506 81

5 CL32538Contig1 50.098 950 59 452


Sorbitol dehydrogenase-like

Crassostrea gigas 534 73

Aplysia californica 530 76

Biomphalaria glabrata 517 74


41

The same affinity chromatography fraction (10 mM NAD+) was then processed by

ion exchange chromatography, using a higher performance column (Mono Q 4.6/100

PE). In this analysis, several protein peaks were observed between fractions 11 and 25

(Figure 10). Therefore, the activity of polyol dehydrogenases was quantified in these

fractions and the protein profile analysed by SDS-PAGE (Figure 11). Once again, two

peaks of activity were detected, the first one with slightly higher activity for mannitol than

sorbitol around fraction 15 and 16, a second peak with an activity for sorbitol two-fold

higher than mannitol around fraction 22.

Seven protein bands of interest, detected after SDS-PAGE of these fractions, were

analysed by MS/MS and further identified (Table 8), using a Patella vulgata database

and by performing a Blastp of the protein at the NCBI database. Bands 1 and 2, from

fractions 16 and 20, respectively, were both identified as being a class III alcohol

dehydrogenase. Band 3 from fraction 22 was identified as a universal stress protein.

Bands 4 and 5, from fraction 15, were both identified as D-arabinitol dehydrogenase 1.

However, this last identified protein was present in low quantity in the samples.

Therefore, 25 of the most abundant peptide ions of each sample were analysed and

glyceraldehyde-3-phosphate dehydrogenase was identified with statistical significance.

Finally, bands 6 and 7, from fraction 22, were identified as sorbitol dehydrogenase-like

protein.


42

Figure 10 - Ion exchange chromatogram of 10 mM NAD+ fraction from affinity chromatography with Mono Q 4.6/100 PE column Proteins eluted with an ascending and continuous NaCl gradient (0.05

M to 1 M), at a flow rate of 1 ml per sec. (green) protein quantification (abs 280 nm); (black) high salt solution concentration; (red) conductivity.


43

Figure 11 - Characterization of the fractions obtained by ion exchange chromatography (Mono Q 4.6/100 PE). Top: analysis of polyol dehydrogenase activity. Bottom: analysis of proteins by SDS-

PAGE. Proteins were stained with Coomassie blue. Molecular weight markers are shown on the left and right. Spots 1-7 were identified by MS/MS: 1 & 2 alcohol dehydrogenase class III; 3 – universal

stress protein; 4 & 5 - D-arabinitol dehydrogenase and Glyceraldehyde-3-Phosphate dehydrogenase 1; 6 & 7 – sorbitol dehydrogenase-like protein (Table 8).


44

Table 8 - Proteins from ion exchange chromatography fraction with Mono Q 4.6/100 PE column (Figure 11) analysed by mass spectrometry (MS/MS) and identified with a Patella vulgata database.

Mascot search result scores greater than 65 are significant (p<0.05). Data shows the highest score identifications for each sample.

Mascot Search NCBI Blastp

Band Accession MW

(kDa) Score

No. Peptides Identified

No. Amino Acids

Specific hits Organism Score Identity

(%)

1

CL24363Contig1 48.891

169 35

390


Alcohol dehydrogenase class III


2 207 41 Aplysia californica 624 80


3 CL27250Contig1 44.983 88 19 180


Universal stress protein Biomphalaria glabrata 230 70

Octopus bimaculoides 221 60

4

CL9544Contig1 44.491

176 29

399

Hypothetical protein LOTGIDRAFT_188066

Lottia gigantea

452 60


5 153 29


D-arabinitol dehydrogenase 1 Neodiprion lecontei 308 45

Ciona intestinalis 303 44

6

CL32538Contig1 50.098

110 17

452


Sorbitol dehydrogenase-like


7 88 14 Aplysia californica 530 76


Table 9 - Peptide ions from ion exchange chromatography fraction with Mono Q 4.6/100 PE column (Figure 11) analysed by mass spectrometry (MS/MS) and identified with UniProt (Metazoa) database.

Mascot search result scores greater than 82 are significant (p<0.05); Data shows the highest score identifications for each sample.

Mascot Search

Band Accession MW (kDa) Score Description

4 C7SP37 20.658 228 Glyceraldehyde-3-Phosphate dehydrogenase (Fragment)

5 G9BBW6 16.599 102 Glyceraldehyde-3-Phosphate dehydrogenase (Fragment)


45

4. Discussion

The capacity to metabolize polyols seems to be an important factor that allows

organisms to deal with stress-inducing conditions through osmoregulation [32, 118], but

also to use them as carbon and energy sources after food intake [24]. Our results using

total homogenates of different gastropod species, showed that almost every species

present dehydrogenase activity for both mannitol and sorbitol, suggesting that this

metabolic capacity is generalized throughout all the gastropod clades. In contrast, the

oxidase activity towards polyols was only detected in herbivorous species of

Caenogastropoda and Heterobranchia clades (data not published).

When correlating polyol dehydrogenase with their diet, it was observed that all

herbivores species possess this activity at least for one of the substrates. On the other

hand, half of the analysed carnivorous species did not show polyol dehydrogenase

activity, and those who did presented dehydrogenase activities to these substrates, were

quite lower levels than the observed in herbivores. These data suggest that the

herbivorous species, due their higher polyol intake with the vegetal matter of the diet,

would have an extra need to metabolize these compounds and therefore higher activity

levels. However, this capacity does not seem to be exclusive for herbivores, since some

carnivores also have it. It is unclear whether this is a conserved feature in these animals,

or an adaptation for those whose diet includes polyols and, therefore, use these

compounds for energy intake and survival.

Following the principle that the metabolic activities detected in gastropods

digestive gland would be tightly correlated with the compounds mainly ingested in the

diet [24], the metabolic capacity of the digestive gland for cinnamyl alcohol was also

studied. This compound is representative of the aromatic alcohols [33, 34]. Since it was

already described its correlation with lignin degradation and its oxidative process being,

in part, similar between fungi and gastropods (physico-chemical parameters) [34],

significant activities for this substrate were expected at least in herbivorous species that

might ingest some lignified plant matter.

Interestingly, a previous study reported cinnamyl alcohol oxidase activity in all the

studied species, herbivores and carnivores, except in O. celtica (data not published). In

this work, the dehydrogenase activity for cinnamyl alcohol was detected in all the 26-

studied species, and for the first time in animals. These results suggest that cinnamyl

alcohol metabolism in gastropod digestive gland in generalized among all the studied

species, without any significant correlation with taxonomy and dietary habits. However,


46

a slight correlation could be established regarding the species habitat, since the studied

freshwater species presented lower cinnamyl alcohol dehydrogenase activity values in

an overall comparison. Therefore, due to the presence of aromatic alcohol metabolism

in carnivores but also in herbivores that do not feed mainly on lignified plants, the

correlation of this metabolism and the consumption of lignified plant matter does not

seem valid anymore. Another hypothesis for this generalized metabolic capacity through

all the analysed species could be some correlation with detoxification processes.

At this point, data suggest that the polyols metabolism is much more correlated

with dietary habits, and perhaps the taxonomy, in the studied species of gastropods,

while aromatic alcohols metabolisms seems to be a much more generalized feature of

these animals.

There is an ongoing discussion about whereas the natural exposure to ethanol is

responsible for the diversity of oxidoreductases [38]. Studies with cephalopods

dehydrogenases showed that only class III ADH is present in these animals [110]. This

data does not support the hypothesis of ethanol metabolism, typically translated by the

presence of class I ADH, in molluscs. In previous studies, the oxidase activity for ethanol

was evaluated in several gastropod species, where only five of the 26-studied species

presented detectable activity for this compound (data not published). Additionally, in the

present study, ethanol dehydrogenase activity was only detected in two herbivorous

eupulmunates. In both studies, the ethanol-related activities detected presented low

overall values when compared with activity values of other substrates. Therefore, this

data suggests that gastropods digestive gland does not play a major role in ethanol

metabolism (via oxidase or dehydrogenase). It is uncertain if other pathways of ethanol

metabolism exist in molluscs, in the digestive gland or other organs.

Several studies have previously shown that alcohol dehydrogenases [119] and

polyol dehydrogenases [91, 105] are capable of metabolizing, not only the main

substrate responsible for its nomination, but also several other similar substrates. In fact,

they are generally seen as quite unspecific enzymes [38]. In some cases, even higher

activities are found for other substrates [105]. Our results also suggest this unspecific

behaviour of these enzymes. This is supported by the results of in situ activity detection,

where each analysed species presented similar dehydrogenase activity profiles towards

various substrates (polyols and aromatic alcohol). Interestingly, with exception of P.

vulgata, all the other species may have more than one enzyme with dehydrogenase

activity for these substrates. However, it cannot be ruled out the hypothesis that the

second band observed in the analysis of these enzymes corresponds to the same protein

post-translationally modified. Additionally, the well-defined band observed in these gels,


47

might be a complex mixture of proteins in which only some of them possess

dehydrogenase activity for these substrates.

The comparison of the relative migration distance for both polyols and aromatic

alcohol dehydrogenases after in situ activity detection suggests that proteins with high

polyols dehydrogenase activity are different from those with high aromatic alcohol

dehydrogenase activity. However, being already known the unspecific behaviour of the

oxidoreductase enzyme family [38, 41, 109], a feature already suggested in this study, it

is not possible to exclude the hypothesis that polyol and aromatic alcohol metabolism

enzymes (oxidases and dehydrogenases) might non-specifically use the substrates of

each other, but with lower efficiency.

Aiming to isolate and identify the proteins with mannitol or sorbitol dehydrogenase

activity, we used cytosolic fractions from Patella vulgata since this species exhibited high

activity values, it is easier to obtain high amounts of tissue due to a big proportion of

digestive gland present in this animal, and this species can be easily collected all year.

The separation of proteins by affinity chromatography allowed the obtaining of

fractions significantly enriched (around ten-fold) in polyols or cinnamyl alcohol

dehydrogenases. Regarding the protein behaviour in this column, this first

chromatography also supported the hypothesis of the significant biochemical

differences, between the proteins responsible for polyols and aromatic alcohol

dehydrogenase activities. Proteins with polyol dehydrogenase activity were eluted from

the column in the initial washing steps, but mainly in the lower NAD+ concentration steps,

and proteins with aromatic alcohol dehydrogenase activity were eluted in the higher

NAD+ concentration steps and in the final washing steps. This protein behaviour

suggests that proteins with polyols dehydrogenase activity present much less affinity for

this column than proteins with aromatic alcohol dehydrogenase activity. In this work, we

focused our attention in the fraction enriched for polyols dehydrogenase activity. Future

studies will be performed to identify cinnamyl alcohol dehydrogenases.

Overall the results show that the Blue Sepharose CL-6B column was quite effective

in the separation of dehydrogenase enzymes, since the characterization of the fractions

enriched in dehydrogenases led to the identification of two dehydrogenases (cytosolic

malate dehydrogenase and GAPDH), one class III alcohol dehydrogenase, and two

polyol dehydrogenases (D-arabinitol dehydrogenase 1 and sorbitol dehydrogenase).

The analysis of dehydrogenase activity in the fractions obtained by ion exchange

chromatography revealed 2 peaks, a first one early eluted from the column that had

mannitol dehydrogenase activity higher than sorbitol dehydrogenase activity, and a

second peak where the sorbitol dehydrogenase activity was higher than mannitol

dehydrogenase activity. These results suggest that different proteins are associated with


48

these dehydrogenases (Figure 11). Consistent with the fact that these type of enzymes

can be quite unspecific [91], they can metabolize both D-mannitol and D-sorbitol, it was

identified a D-arabinitol dehydrogenase (ARDH) in the first peak, with higher

dehydrogenase activity for mannitol (band 3 of Figure 9 and band 4 and 5 of Figure 11),

and a sorbitol dehydrogenase-like protein, with higher dehydrogenase activity for sorbitol

(band 5 of Figure 9 and bands 6 and 7 of Figure 11).

D- and L-arabinitol dehydrogenases are enzymes mostly studied in fungi [120,

121]. Interestingly, despite having an apparent classification as a polyol dehydrogenase,

there are some evidences that this enzyme has functions similar to that of medium-chain

dehydrogenases [122] or even short- and long-chain dehydrogenases [123, 124]. Thus,

our Blastp results identified the founded protein as from MDR superfamily and higher

scores were obtained for three uncharacterized sequences of Lottia gigantea, the insect

Neodiprion lecontei and cordate Ciona intestinalis. This data reinforced the idea that little

is known about arabinitol dehydrogenases in other organisms than fungi. However, this

might be one of the first evidences of this enzyme in molluscs.

Regarding the overall data of this work, especially the results related with polyols

dehydrogenases, were obtained the first clues on which enzyme may be responsible for

each dehydrogenase activity. However, more work must be done for the purification of

these enzymes, in order to characterize the enzymes with polyols dehydrogenase

activity. Additionally, more work must be focused in the proteins with aromatic

dehydrogenase activity.


49

5. Conclusions and perspectives

With this study, it was revealed for the first time the presence of dehydrogenase of

mannitol and aromatic alcohols in the digestive gland of gastropods. It was reinforced

the biochemical differences between proteins of polyol and aromatic alcohols

metabolisms. Also, this study supports the absence of enzymes with significant ethanol

dehydrogenase activity in molluscs, having the gastropods digestive gland by example.

For a better idea of which specific ADH classes are present in these organisms,

biochemical assays, regarding that ADH class specificity, could be done in the future

studies.

The first steps for isolation of aromatic alcohol and polyalcohol dehydrogenases in

the marine gastropod Patella vulgata, resulted in the identification of at least two

enzymes with polyol dehydrogenase activity (sorbitol and arabinitol dehydrogenases). A

similar line of research regarding the proteins with aromatic alcohols dehydrogenase

activity, namely cinnamyl alcohol, is yet to be developed in future studies. More studies

in the future could be focused in arabinitol dehydrogenase because of the lack of

knowledge about this enzyme in organisms other than fungi.

Thus, with these new knowledge’s, there is a changing of perspectives about the

metabolisms of gastropod`s digestive gland, but also a comparison point for this type of

enzymes with other organisms.

Finally, future work aims to identify the genes codifying these proteins, for their

subsequent heterologous production and to study their metabolic profiles recurring to

other alcoholic substrates.


50

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Annex


58

Table S1 - Oxidase activities in the digestive gland of gastropods (nmol.min-1. g-1 ± SD). Shaded lines correspond to carnivorous species (data not published).

Species Mannitol Sorbitol Cinnamyl alcohol Ethanol

Patellogastropoda Patella vulgata - - 283.4 ± 168.8 95.4 ± 44.2

Neritimorpha Neritina natalensis - - 35.8 ± 14.8 -

Vetigastropoda

Phorcus lineatus - - 120.0 ± 53.9 -

Gibbula umbilicalis - - 87.3 ± 41.7 -

Calliostoma zizyphinum - - 241.6 ± 76.1 -

Caenogastropoda

Nassarius reticulatus - - 713.7 ± 110.8 -

Charonia lampas - - 156.2 ± 72.3 41.1 ± 18.6

Nucella lapillus - - 223.6 ± 84.5 -

Hexaplex trunculus - - 290.0 ± 96.8 -

Ocenebra erinaceus - - 319.8 ± 51.2 -

Littorina littorea 472.6 ± 147.8 - 609.2 ± 208.2 -

Pomacea bridgesii 203.3 ± 58.8 17.6 ± 14.5 796.0 ± 81.5 -

Marisa cornuarietis 217.3 ± 138.4 6.6 ± 7.8 692.3 ± 146.0 -

Heterobranchia

Pleurobranchaea meckelii - - 174.8 ± 71.0 -

Doriopsilla areolata - - 94.2 ± 25.7 -

Armina maculata - - 353.2 ± 380.0 -

Aglaja tricolorata - - 259.7 ± 93.3 -

Haminoea orbignyana 329.9 ± 233.6 - 1,098.8 ± 685.9 -

Bulla striata 126.1 ± 63.6 - 244.4 ± 102.3 -

Aplysia depilans 651.7 ± 107.2 - 889.5 ± 245.5 -

Siphonaria pectinata 238.5 ± 98.5 - 506.9 ± 211.7 -

Planorbarius corneus 68.2 ± 41.7 - 450.1 ± 205.1 113.3 ± 47.6

Onchidella celtica 1,462.3 ± 246.9 122.8 ± 48.1 - -

Cornu aspersum 720.0 ± 106.8 25.9 ± 44.4 476.3 ± 304.6 395.2 ± 141.2

Arion ater 1,716.1 ± 868.4 70.4 ± 29.9 2,508.2 ± 628.8 642.4 ± 229.3

Lehmannia valentiana 1,417.3 ± 454.7 90.4 ± 58.3 103.5 ± 61.9 48.5 ± 30.8

and polyol dehydrogenases in patella vulgata and · grupos de organismos. ... o segundo maior filo...

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