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Revista de Biología Tropical, ISSN: 2215-2075, Vol. 71: e51085, enero-diciembre 2023 (Publicado Mar. 02, 2023)
Ontogeny of digestive enzymes in larvae of the Clown Anemonefish,
Amphiprion ocellaris (Perciformes: Pomacentridae)
Gabriela Velasco-Blanco1; https://orcid.org/0000-0002-0288-8141
Carlos Alfonso Álvarez González2; https://orcid.org/0000-0001-9240-0041
Maria Isabel Abdo de la Parra1; https://orcid.org/0000-0003-2148-9661
Luz Estela Rodríguez-Ibarra1; https://orcid.org/0000-0001-8369-1000
Leonardo Ibarra-Castro3; https://orcid.org/0000-0002-2159-9038
Claudia I. Maytorena-Verdugo4; https://orcid.org/0000-0001-7417-858X
José Natividad Arias-Jiménez2; https://orcid.org/0000-0001-6447-3112
Emyr Saul Peña Marín5*; https://orcid.org/0000-0002-4736-9089
1. Centro de Investigación en Alimentación y Desarrollo A.C., Mazatlán Unit., Av. Sábalo Cerritos s/n, Mazatlán, Sinaloa
89010, México; gvelas@ciad.mx, abdo@ciad.mx, eibarra@ciad.mx
2. Laboratorio de Fisiología en Recursos Acuáticos, División Académica de Ciencias Biológicas, Universidad Juárez
Autónoma de Tabasco, Km 0.5 carretera Villahermosa- Cárdenas entronque Bosques de Saloya, Villahermosa,
Tabasco, México; alvarez_alfonso@hotmail.com, aijn930427@gmail.com
3. Program in Fisheries and Aquatic Sciences, School of Forest, Fisheries, and Geomatics Sciences, Institute of Food
and Agricultural Sciences, University of Florida, 7922 Northwest 71st Street, Post Office Box 110600, Gainesville,
Florida 32611, USA; and Whitney Laboratory for Marine Bioscience; 9505 Oceanshore Blvd, St Augustine, Florida,
32080, USA.; l.ibarracastro@ufl.edu
4. Universidad Politécnica del Centro, Carretera Federal, Villahermosa-Teapa Km 22.5, Tumbulushal Centro, 86290,
Villahermosa, Tabasco, México; clau.maytorena@gmail.com
5. Instituto de Investigaciones Oceanológicas, Universidad Autónoma de Baja California (UABC), Ensenada 21100,
México; emyr.pea@uabc.edu.mx, ocemyr@yahoo.com.mx (*Correspondance)
Received 25-X-2022. Corrected 01-XII-2022. Accepted 23-II-2023.
ABSTRACT
Introduction: The Clown anemonefish (Amphiprion ocellaris) is the most popular fish species in the marine
aquarium trade; however, there is a lack of information on their digestive physiology during larval ontogeny,
valuable information needed for diet design and management protocols.
Objective: To characterize the early digestive enzymes of A. ocellaris larvae.
Methods: We used three pools (10 larvae each) and extracted 10 samples per tank, from just before hatching
to the 38th day. We analyzed the specific activity of acid and alkaline proteases, trypsin, chymotrypsin, leucine
aminopeptidase and lipase; and did acid and alkaline protease zymograms.
Results: We detected all measured enzymes at hatching. Acid proteases increased in activity until the 38th day.
Alkaline proteases, trypsin, chymotrypsin, and leucine aminopeptidase had the same pattern, and maximum
activity on the 8th day, decreasing at the 38th day. Lipase activity peaked on the 8th and 30th day. The acid
zymogram had a single band, appearing on the 8th day. A total of eight alkaline proteases were revealed (154.2,
128.1, 104.0, 59.8, 53.5, 41.9, 36.5 and 25.1 KDa), with seven bands on the 1st day and all bands from the 3rd
to 8th day, decreasing at two bands (41.9 and 25.1 KDa) in the 38th day.
https://doi.org/10.15517/rev.biol.trop..v71i1.51085
BIOTECHNOLOGY
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INTRODUCTION
Clown anemonefish (Amphiprion ocel-
laris) belongs to the subfamily Amphiprioninae
(family Pomacentridae) with 28 species and
are the most popular and commercial exploited
species of fishes in the marine aquarium orna-
mental trade (Khoo et al., 2018; Khoo et al.,
2019). These species are considered as a model
for scientific research, especially in nutritional
studies, initial egg ontogeny and the quality of
larvae (Delbare et al., 1995). Several studies
in A. ocellaris report the reproductive biology,
early development, including embryology and
larval stage, osteological development of the
vertebral column and caudal complex, growth
pattern and natural diet (Khoo et al., 2018;
Khoo et al., 2019; Madhu et al., 2012; Rodrí-
guez-Ibarra et al., 2017; Yasir & Qin, 2007).
Even embryo development organogen-
esis information is available in the species.
Nevertheless, larval ontogeny includes the
transition between endogenous to exogenous
energetic reserves, a period considered as a
bottleneck in ornamental aquaculture spe-
cies (Olivotto et al., 2011), where several
morpho-physiological processes take place,
including digestive enzyme development (Chen
et al., 2019; Wilson & Castro, 2010; Zam-
bonino-Infante & Cahu, 2001). In this sense,
nutrient availability for metabolic functions is
directly influenced by the enzyme digestion
process, which depends mainly on the type of
digestive enzymes (Nazemroaya et al., 2015).
The development of specific feeds for larvae
requires a detailed knowledge of their digestive
physiology because of many marine fish larvae
show null or a rudimentary digestive system at
hatching and during digestive development, the
digestion, absorption, and nutritional require-
ments are in constant change (Rønnestad et
al., 2013; Shin et al., 2022). In addition, the
digestive system development varies in time,
depending on the ontogeny type (direct, tran-
sitional, or indirect) of each species (Balon,
1986; Balon, 1990; Peñaz, 2001), where many
marine fish larvae lack a functional stomach
and they depend on pancreatic proteases for
protein digestion (Rønnestad et al., 2013).
Thus, an understanding of the changes in the
ability to digest and absorb during each larval
stage is needed to improve productivity.
Conclusion: A. ocellaris has a functional stomach on the 8th day, and, on the 38th day, a digestive omnivore
pattern with a tendency to carnivory.
Key words: digestive development; early ontogeny; electrophoresis; larvae; lipases; proteases.
RESUMEN
Ontogenia de enzimas digestivas en larvas del pez payaso,
Amphiprion ocellaris (Perciformes: Pomacentridae)
Introducción: El pez payaso (Amphiprion ocellaris) es la especie de pez más popular en el comercio de acuarios
marinos; sin embargo, falta información sobre su fisiología digestiva durante la ontogenia larval, información
valiosa necesaria para protocolos de diseño y manejo dietético.
Objetivo: Caracterizar las enzimas digestivas tempranas de larvas de A. ocellaris.
Métodos: Usamos tres homogenados (con 10 larvas cada uno) y extrajimos 10 muestras por tanque, justo antes
de la eclosión hasta el día 38. Analizamos la actividad específica de proteasas ácidas y alcalinas, tripsina, quimo-
tripsina, leucina aminopeptidasa y lipasa; e hicimos zimogramas de proteasas ácidas y alcalinas.
Resultados: Detectamos todas las enzimas medidas en la eclosión. La actividad de proteasas ácidas incrementó
hasta el día 38. Proteasas alcalinas, tripsina, quimotripsina, y leucina aminopeptidasa tuvieron el mismo patrón,
con actividad máxima en el octavo día, decreciendo en el día 38. Hubo picos en la actividad lipasa a los ocho y 30
días. El zimograma ácido tuvo una banda única, apareciendo al octavo día. Se hallaron ocho proteasas alcalinas
(154.2, 128.1, 104.0, 59.8, 53.5, 41.9, 36.5 y 25.1 KDa), con siete bandas al primer día, y todas las bandas entre
el tercer y octavo día, bajando a dos bandas (41.9 y 25.1 KDa) al día 38.
Conclusión: A. ocellaris tiene un estómago funcional al octavo día, y, al día 38, un patrón digestivo omnívoro
con tendencias carnívoras.
Palabras clave: desarrollo digestivo; electroforesis; larvas; lipasas; ontogenia temprana; proteasas.
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Revista de Biología Tropical, ISSN: 2215-2075, Vol. 71: e51085, enero-diciembre 2023 (Publicado Mar. 02, 2023)
Yúfera et al. (2018) proposed the classifi-
cation of enzymes during ontogeny by the pat-
tern of appearance, using normalized data on
degree days and the maximum activity detected
in each enzyme type. Therefore, enzymes could
be classified as precocious, medium, or late
enzyme development. According to previous
studies in A. ocellaris, at hatching, larvae pos-
sess an advance development on their diges-
tive system (Madhu et al., 2012; Yasir & Qin,
2007). Nevertheless, by a short period time,
larvae show embryo vestiges such as yolk-sac,
and is still developing several systems, for this
reason A. ocellaris, as well to other Amphiprion
species, shows a transitional early ontogeny
development, characterized by a long embry-
onic period before hatching and short larvae
period before juvenile transforming.
Therefore, larvae culture requires the char-
acterization of digestive system changes during
larvae development to understand specific spe-
cies adaptations and generate specific feeding
protocols and development of specific feeds.
Many studies are focus to characterize the
development of the digestive system during
the early ontogeny, including golden pompano
(Trachinotus ovatus) (Ma et al., 2014), sobaity
sea bream (Sparidentex hasta) (Nazemroaya
et al., 2015), black tetra (Gymnocorymbus ter-
netzi) (Lipscomb et al., 2020), red sea bream
(Pagrus major) (Khoa et al., 2019), japanese
flounder (Paralichthys olivaceus) (Khoa et al.,
2021), japanese eel (Anguilla japonica) (Shin
et al., 2022). However, to our knowledge, only
one report on digestive enzymes in A. ocellaris
exists, showing the gastric emptying and pepsin
activity on juvenile stage (Khoo et al., 2019).
Despite the economic importance of clown
anemonefish commercialization, information
on the enzymatic profile and digestive capacity
of the specie is scarce. In addition, the great
diversity, and variants of the development
between species made necessary to character-
ize each species. Hence, the aim of the pres-
ent study was to characterize the ontogeny of
digestive proteases and lipase in clown anem-
onefish larvae (A. ocellaris).
MATERIALS AND METHODS
Larvae rearing: Fertilized eggs were
obtained from an A. ocellaris couple spawned
in Centro de Investigación en Alimentación y
Desarrollo A.C. (CIAD), Mazatlán. Eggs were
transferred to three 300 l water volume tanks
for presumptive density of 1.5 larvae/l. Tem-
perature was maintained at 26 ± 1.0 °C, salinity
of 35 ± 1.0 g/l and a dissolved oxygen concen-
tration of 6.2 mg/l. Tanks had constant aeration
(1-2 l/min) with 30 to 40% water exchanges
every third day by tank siphoning until day
13. From day 14, continuous water flow was
adjusted to start with two total changes per day,
reaching four total water changes at the end of
the experiment. The larvae total length (TL)
was measured (± 0.01 mm) with a binocular
microscope and larvae growth was determined
as the total growth rate (TGR) as mm/day: TGR
= (final TL – initial TL) /days.
The feeding protocol is shown in Table 1.
Briefly, larvae were reared using the green water
technique from the 1st to 6th day after hatching
(DAH) using microalgae Nannochloropsis ocu-
lata, from the 2nd to 9th DAH rotifers of the
species Brachionus rotundiformis were added.
From the 7th to 16th DAH, nauplii of Artemia
sp. were added, while from the 14th to 38th
DAH, a formulated diet (Skretting) was used.
During early development, 10 samples per tank
of egg just before hatching (day 0) and larvae
at 1, 3, 5, 8, 14, 20, 28 and 38 DAH were
taken in 1.5 ml Eppendorf tubes and frozen
at -80 °C for two days, lyophilized and stored
in dry conditions at -20 °C until analysis. For
comparisons between closely related species,
thermal age units (cumulative degree-DAH,
CTU) were used.
Preparation of enzymatic extracts:
Three larvae pool (10 larvae each pool) were
homogenized separately in a 1:100 (weight:
saline solution 0.9 % NaCl) ratio using an Ultra
Turrax (IKA T18 basic, Wilmington, USA).
The homogenates were centrifuged (14 000 g
for 15 min at 4 °C) and the supernatants were
recovered and stored (-80 °C) until analysis.
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The Bradford (1976) technique was used to
determine soluble protein concentration in the
larvae enzymatic extracts at each stage.
Digestive enzyme activity analysis: Acid
proteases activity was determined using bovine
hemoglobin 1 % as substrate in 100 mmol/l
glycine-HCl buffer at pH = 2 (Anson, 1938).
Alkaline proteases activity was determined
using casein 1 % as substrate in 100 mmol/l
Tris-HCl + 10 mmol/l CaCl2 buffer at pH
= 9 (Walter, 1984). The trypsin activity was
determined using Nα-Benzoyl-DL-arginine-4-
nitroanilide hydrochloride (BAPNA) 1 mmol/l
as substrate in 100 mmol/l Tris-HCl + 10
mmol/l CaCl2 buffer at pH = 8 (Erlanger et al.,
1961). Chymotrypsin activity was quantified
using SAAPNA (N-succinyl-ala-ala-pro-phe
p-nitroanilide) 0.1 mmol/l as substrate in 100
mmol/l Tris-HCl solution + 10 mmol/l CaCl2
buffer at pH = 7.8 (Del-Mar et al., 1979). The
leucine-aminopeptidase activity was measured
using leucine p-nitroanilide 1 mmol/l as sub-
strate in 50 mmol/l monobasic sodium phos-
phate buffer at pH = 7.2 (Maroux et al., 1973).
The lipase activity was measured using
4-Nitrophenyl acetate 200 mmol/l as substrate
in buffer 50 mmol/l Tris-HCl at pH = 7.2
with sodium taurocholate 100 mmol/l. Activ-
ity revelation was performed using Fast blue
solution (100 mmol/l) and was clarify by add-
ing ethanol: ethyl acetate solution (1:1 v/v)
(Versaw et al., 1989).
All enzymatic activities were performed at
37 °C by triplicated and products were quantified
in a Genesys 10S UV-Vis spectrophotometer
(ThermoScientific, Waltham, MA, USA). The
tyrosine released from hemoglobin and casein
hydrolysis was determined at 280 nm, the
amount of p-nitroanilide released from BAPNA,
SAAPNA and L-leucine-p-nitroanilide hydro-
lysis was determined at 410 nm and the amount
of p-nitrophenol released from 4-Nitrophenyl
acetate hydrolysis was determined at 405 nm.
Total activity (Units/ml) = [Δabs*reaction
final volume (ml)] / [MEC*time (min)*extract
volume (ml)] and Specific activity (Units/mg
prot) = Total activity / soluble protein (mg/ml),
where Δabs represent the increase in absor-
bance, and MEC represents the molar extinc-
tion coefficient of tyrosine, p-nitroaniline and
p-nitrophenol (0.005, 0.008 and 0.02 ml/µM/
cm, respectively).
Native and SDS-PAGE zymogram: Mini-
PROTEAN 3 Cell (Bio-Rad) with four vertical
plate gels (8 × 10 × 0.075 cm) with 10 ml
sample capacity per plate was used for the elec-
trophoretic analyses.
Acid proteases electrophoresis was run
under non-denaturing native conditions (Native-
PAGE) with continuous polyacrylamide (PAA)
(10 %) in Tris (25 mmol/l) and glycine buffers
(192 mmol/l, pH = 8.3, 80 volts) according to
Davis (1964). Alkaline proteases electropho-
resis was run under denaturalizing conditions
(SDS-PAGE), with PAA stacking a gel (4 %)
and PPA resolving gel (10 %), adding SDS
(0.1 %) in Tris buffer (25 mmol/l) and glycine
(192 mmol/l, pH = 8.3, 100 volts), according
to Laemmli (1970) and adapted by García-
Carreño et al. (1993).
Table 1
Feeding schedule and food types during clown anemonefish (Amphiprion ocellaris) larvae rearing
Food item Type Amount Period (DAH)
Microalgae Nannochloropsis oculata 500 000 cel/ml 1-6
Rotifer Brachionus rotundiformis 10 rotifer/ml 2-6
Rotifer Brachionus rotundiformis 5 rotifer/ml 7-9
Artemia Artemia sp. nauplii 6 nauplii/ml 7-13
Artemia Artemia sp. nauplii 3 nauplii/ml 14-16
Dry microdiet Formulated diet (Scretting) At libitum 14-38
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Native-PAGE electrophoresis gels were
revealed for proteases isoforms according to
the procedure of Díaz-López et al. (1998). The
removed gels were soaked during 15 min in
100 mmol/l Tris-HCl in pH = 2.0 to activate
enzymes. Then, gels were submerged during 60
min at 25 °C in 0.25 % hemoglobin solution in
Glycine-HCl buffer (100 mmol/l) at pH = 2.0.
The gels were washed with distilled water and
fixed in 12 % trichloroacetic acid (TCA) solu-
tion for 15 min. SDS-PAGE electrophoresis
gels were washed and incubated during 60 min
at 4 °C in a 0.5 % casein solution (Tris-HCl 100
mmol/l buffer, pH = 9). Then, gels were incu-
bated for 60 min at 37 °C in the same solution,
and then washed and fixed in TCA as previ-
ously described. For Native and SDS-PAGE,
gels were stained using a 0.1 % Coomassie bril-
liant blue R-250 solution, while distaining was
carried out in a 35:10:55 solution of methanol-
acetic acid-water (Weber & Osborn, 1969).
Clear zones revealed the activity of proteases,
the bleaching process was maintained until
well-defined zones were obtained (after 2-4 h).
The protein leader (Termo Scientific; Cat#
26614) comprised of 14 recombinant proteins
(weighting from 10 kDa to 200 kDa) was used
as molecular markers and 5 µl per well was
applied to each SDS-PAGE electrophoresis.
Molecular weight (MW) of each band was cal-
culated using a linearly adjusted relationship
between the Rf (migration distance of the pro-
tein/ migration distance of the dye front) and
log10 of the MW of each recombinant protein.
Statistical analysis: The specific activ-
ity (U/mg protein) was analyzed for normality
(KS) and homoscedasticity (Levene) postulates
and one-way analysis of variance (ANOVA)
was applied to determine if differences exist
between age and enzyme activities. When dif-
ferences exist between treatments, a posteriori
Tukey test was used. All tests were carried out
using a level of significance of 0.05. The Sigma
Plot (Systat Software inc., 2009) was used to
perform the statistical analyzes.
RESULTS
Growth performance: At hatching, A.
ocellatus larvae showed a total length of 4.5 ±
0.13 mm, increasing to 14.73 ± 1.01 mm at 38
DAH, displaying a TGR value of 0.2 mm day-1.
Growth of A. ocellatus larvae fit to an exponen-
tial model (r2 = 0.9506) (Fig. 1).
Fig. 1. Larvae growth curve in total length (mean ± SD)
of clown anemonefish (A. ocellaris) from hatching to 38
DHA.
Enzyme activity: For all enzymes, activ-
ity was detected from the moment of hatch-
ing (1 DAH), with values of 0.150, 0.416,
0.167, 41.65, 0.812 and 10.01 U/mg protein for
acid proteases, alkaline proteases, trypsin, chy-
motrypsin, LAP, and lipase respectively. The
specific activity (U/mg protein) and relative
activity (%) against degree days of acid pro-
teases shows two peaks at day five (130 CTU)
and day 28 (728 CTU), decreasing to day 38
(988 CTU) (P ≤ 0.05), while total alkaline
proteases show a peak at day 8 (208 CTU) and
then decreased from day 14 (364 CTU) to day
38 (988 CTU) (P ≤ 0.05) (Fig. 2A, Fig. 2B).
The trypsin, chymotrypsin and leucin amino-
peptidase activity shows the same pattern in
specific activity (U/mg protein) and relative
activity (%) against degree days, it increased
from hatching with a peak at day 8 (208 CTU)
and then decreased from day 14 (364 CTU)
to day 38 (988 CTU) (P ≤ 0.05) (Fig. 2C, Fig.
2D). The lipase specific activity (U/mg protein)
6Revista de Biología Tropical, ISSN: 2215-2075 Vol. 71: e51085, enero-diciembre 2023 (Publicado Mar. 02, 2023)
and relative activity (%) against degree days
showed two higher peaks at day 8 (208 CTU)
and day 28 (728 CTU), decreasing at day 38
(988 CTU) (P ≤ 0.05) (Fig. 2E, Fig. 2F).
Electrophoretic analyzes: Acid zymo-
gram shows a single band with acid activity
during all ontogeny, which appears at the 8th
DAH and increases its activity until the 38th
DAH (Fig. 3A). Alkaline zymogram shows
a total of eight bands (154.2, 128.1, 104.0,
59.8, 53.5, 41.9, 36.5 and 25.1 KDa) (Fig. 3B).
At 0 DAH, no bands were detected. On the
1st DAH (26 CTU) the heaviest seven bands
Fig. 2. Specific enzyme activity (U mg protein-1) and relative activity (%) in larvae development of clown anemonefish
(A. ocellaris) (mean ± SD). Statistical differences (P < 0.05) against stages DAH are indicated by superscript letters. A.
Specific activity of acid and alkaline proteases; B. Relative activity of acid and alkaline proteases against degree days; C.
Specific activity of trypsin, chymotrypsin, and leucine-aminopeptidase; D. Relative activity of trypsin, chymotrypsin, and
leucine-aminopeptidases against degree days; E. Specific activity of lipase; F. Relative activity of lipase against degree days.
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were revealed, from the 3rd (78 CTU) to 8th
DAH (208 CTU), eight bands were revealed.
At the14th DAH (364 CTU), three bands were
revealed (41.9, 36.5 and 25.1 KDa), at the 20th
DAH (520 CTU) four bands were revealed
(154.2, 128.1, 41.9 and 25.1 KDa), while from
the 28th (728 CTU) to 38th DAH (988 CTU)
two bands were revealed (41.9 and 25.1 KDa).
DISCUSSION
The sequence apparition of most devel-
opmental events in fishes are mostly stable,
only varying in rate and duration of yolk sac
depletion, that depends on the size of the yolk-
sac (Peñaz, 2001). Newly hatched fish larvae
initially depend on endogenous reserves for
survival, but they must switch to exogenous
feeding before their yolk reserve depletes
(Chen et al., 2019); however, this switch is
variable in time, depending on the development
stage at hatching time that defines the ontog-
eny type of each species (Balon, 1986; Balon,
1990; Peñaz, 2001). In this sense, A. ocellaris
possess an advanced development of the diges-
tive system at hatching (Madhu et al., 2012;
Yasir & Qin, 2007). Nevertheless, by a short
period time, larvae show embryo vestiges such
as yolk-sac, and is still developing several sys-
tems, for this reason A. ocellaris, as well as to
other Amphiprion species, shows a transitional
early ontogeny development, characterized by
a long embryonic period before hatching and
short larvae period before metamorphosing
into a juvenile.
The enzymatic characterization in the pres-
ent study was performed from the embryonic
Fig. 3. View of the electrophoretic gels. A. Native-PAGE; B. SDS-PAGE electrophoresis analysis of clown anemonefish (A.
ocellaris) during larvae ontogeny from hatching to 38 days after hatching (DAH). Molecular Marker (MW) protein leader
(Termo Scientific; Cat# 26614) comprised by 14 recombinant protein weighting from 10 kDa to 200 kDa.
8Revista de Biología Tropical, ISSN: 2215-2075 Vol. 71: e51085, enero-diciembre 2023 (Publicado Mar. 02, 2023)
period (before hatching), the eleutheroembryo
period, from hatching and the last day until yolk
sac is digested (from the 1st to 3rd DAH) and
larvae period, with two phases: Protopterygio
phase (encompasses the interval between tran-
sition to exogenous feeding; from the 4th to 7th
DAH) and Pterygiolarvae phase (the jaws are
well developed into functional structures); and
the juvenile period represents the time when
transition to definitive organs is completed
between the 15th to 17th DAH (Madhu et al.
2012; Yasir & Qin, 2007).
Reports in A. ocellaris shows that newly
hatched larvae show the lower jaw formed, but
the mouth is not yet open, however in a few
hours, the mouth is visible with 170 to 210 µm,
able to detect and ingest exogenous food on the
first day after hatching, related to well-devel-
oped sensory and olfactory systems (Madhu et
al., 2012; Yasir & Qin, 2007). Studies in fire
clownfish (A. melanipus) shows that the gut is
already looped and differentiated (foregut, mid-
gut, and hindgut) with large intestinal lumen
and visible food inside the lumen at hatching,
present compact and granular liver, with hepa-
tocytes lacked vacuoles (Green & McCormick,
2001). In tomato clownfish (A. frenatus) larvae
at 24 h after hatching show an evident alimen-
tary tract with gut and liver differentiated. At
the 2nd DAH, a well-developed alimentary tract
is evident, with a distinct stomach, midgut
and hindgut, liver, and pancreas (Putra et al.,
2012). In orange clownfish (A. percula), the
alimentary tract, liver and pancreas are present
at hatching, showing absorptive and digestive
capabilities, where larvae start exogenous feed-
ing immediately (Gordon & Hetch, 2002; Önal
et al., 2008). Red saddleback anemonefish (A.
ephippium) hatch at day 6 post fertilization and
at that moment show open mouth and start feed-
ing by itself and at the 2nd DAH, larvae show a
well-developed digestive system (Rohini et al.,
2018). In this sense, the present study shows
that A. oceallaris presents acid proteases, alka-
line proteases, cytosolic proteases, and lipases
at hatching, which corroborates the existence
of a functional stomach, pancreas, and intestine
with the capacity to acquire exogenous reserves
from food digestion in combination with the
use of endogenous reserves from the yolk sac.
In the present study, alkaline proteases and
lipase show activity from hatching and increase
from first day until the 8th DAH. Complemen-
tary, seven active digestive alkaline proteases
were detected in the zymogram (154.2, 128.1,
104.0, 59.8, 35.5, 41.9 and 36.5 KDa) from
the 1st DAH, increasing one more active band
(25.1 KDa) until the 8th DAH and decreas-
ing in activity and digestive bands from that
moment, showing only two alkaline proteases
(41.9 and 25.1 KDa) at the 38th DAH. In A.
percula, acidophilic zymogen granules precur-
sors of pancreatic enzymes are accumulated in
exocrine pancreatic cells just before hatching
(Önal et al., 2008; Zambonino-Infante & Cahu,
2001), indicating that digestive machinery is
ready. Therefore, the increasing number of
alkaline proteases until the 8th DAH is related
to rapidly morphophysiological changes such
as expansion of support organs (liver and pan-
creas), increase in mean gut epithelium cell
height (increase with age and size), increase in
surface area for enzyme secretion, digestion of
nutrients and absorption, developed vacuoles
in the intestine as functional areas of lipid and
glycogen storage that are of the extracellular
digestion of lipids, diffusion of fatty acids into
enterocytes and re-synthesis of triglycerides in
to the enterocytes (Green & McCormick, 2001;
Gordon & Hetch, 2002; Zambonino-Infante
& Cahu, 2001).
In gastric fishes, the development of a
functional stomach with pepsin-like enzyme
secretion is associated with the transition from
larvae to juvenile, critical to evaluate the diges-
tive capacity of fish larvae, especially when
considering the weaning time (transition from
live prey to formulated feeds) (Kolkovski,
2001; Salze et al., 2012; Zambonino-Infante &
Cahu, 2007). Reports in A. ocellaris show that
at the 7th DAH, gastric glands are established in
the epithelium of the stomach, being capable of
digesting formulated diets at 10 DAH (Gordon
& Hetch, 2002). In the present study, A. ocel-
laris shows a peak of activity in acid proteases
at 5th DAH, appearing one pepsin-like enzyme
9
Revista de Biología Tropical, ISSN: 2215-2075, Vol. 71: e51085, enero-diciembre 2023 (Publicado Mar. 02, 2023)
band (Rf= 0.72) on the acid zymogram at 8th
DAH, showing an increase in activity and
thickness of bands until 38th DAH, associated
to a rapid stomach growth, secretions increase,
as well as to functionality of time retention
(Khoo et al., 2019; Salze, et al., 2012). Reports
in A. percula shows that gastric glands are
established in the epithelium between the 7th
DAH (Gordon & Hetch, 2002) and 11th DAH
(Önal et al., 2008). In A. melanopus, the stom-
ach is visible as a slight enhancement of the
esophagus at the 3rd DAH, with a sphincter in
each end, and lined with cuboidal epithelium
with many secretory cells (Green & McCor-
mick, 2001). Therefore, the decrease in alkaline
proteases activity and the increase in acid pro-
teases activity showed between the 8th and 14th
DAH, correspond to the turnover to a juvenile
period in A. ocellaris. Laboratory management
of A. ocellaris in CIAD-Mazatlán begins with
a dry diet feeding at the 14th DAH, however,
these results show that early weaning time can
be achieved in the species, where reports in A.
percula shows that can be weaned off from 7th
DAH (Gordon et al., 2000).
A. ocellaris is considered as an omnivorous
species because the stomach content, showing a
variety of larvae and algae (Khoo et al., 2018),
and possess a high support of protein digestive
capacity by the stomach, showing a pepsin
activity peak at 2 h after feeding with long time
food retention (36 h), where the gastric secre-
tions are regulated effectively with infrequent
and irregular meals (Khoo et al., 2019). The
switch of acid to alkaline proteases between the
8th and 14th DAH found in the present study,
support that protein digestion in the species is
mainly produced by the stomach, showing only
two alkaline proteases at the 38th DAH. Even if
A. ocellaris feeds on animal and vegetable pro-
tein in their natural environment, the diversity
of digestive alkaline proteases at juvenile stage
is reduce, therefore could be classified as an
omnivorous species with carnivory tendency.
However, more studies on digestive enzyme
functional characterization, as well to digest-
ibility of different protein and lipid sources by
enzymatic battery of the species is required.
Roux et al. (2021) presented standard
methods to maintain breeding pairs of A. ocel-
laris in captivity to establish regular good qual-
ity spawning, and protocols to ensure larval
survival, with the objective to use the species
as experimental marine model. In this study, a
homemade feed was manufactured for breeders
to ensure nutritional quality, however, larvae
nutrition was established using green micro-
algae, rotifers and Artemia nauplii. Therefore,
the proposed method can be improved by the
understanding of the digestive capacity of the
species, considering breeders and larvae to
design specific diets for the different life stages
of A. ocellaris.
According to Yúfera et al. (2018), A. ocel-
laris shows a precocious enzyme development
on total alkaline proteases, trypsin, chymotryp-
sin, and LAP, peaking at 8th DAH (208 CTU),
while pepsin and lipase could be classified as
late enzyme development, peaking at 28th DAH
(728 CTU). However, lipase showed 98 % of
relative activity (%) at 8th DAH regarding to
maximum activity, therefore showed two major
peaks, and could be considered as precocious.
Therefore, A. ocellaris is a precocious species
in protein digestion mainly supporting alkaline
digestion by pancreatic (trypsin, chymotryp-
sin, and lipase) and intestine (membranous
and cytosolic enzymes) enzymes (Zambonino-
Infante & Cahu, 2007), showing a switch
to stomach digestion by pepsin in the last
ontogenic days.
In conclusion, clown anemonefish (A.
ocellaris) shows pancreatic and luminal active
digestive enzymes at hatching, increasing in
activity and enzyme number until the 8th DAH,
time at which an enzymatic switch by the
increasing activity of one acid protease and
decrease of alkaline proteases in activity and
number, however lipase increased its activity on
juvenile stage. Therefore, the protein digestive
capacity in A. ocellaris is mainly supported by
the stomach, showing an enzymatic pattern of
an omnivore fish with a tendency to carnivory.
Ethical statement: the authors declare
that they all agree with this publication and
10 Revista de Biología Tropical, ISSN: 2215-2075 Vol. 71: e51085, enero-diciembre 2023 (Publicado Mar. 02, 2023)
made significant contributions; that there is no
conflict of interest of any kind; and that we fol-
lowed all pertinent ethical and legal procedures
and requirements. All financial sources are
fully and clearly stated in the acknowledge-
ments section. A signed document has been
filed in the journal archives.
All procedures of fish manipulation,
including the euthanasia method by thermal
shock followed the Mexican legislation accord-
ing to Norma Oficial Mexicana NOM-062-
ZOO-1999 from Secretaría de Agricultura,
Ganadería, Desarrollo Rural, Pesca y Aliment-
ación (SAGARPA), the Mexican standards for
good welfare practices of laboratory animals.
ACKNOWLEDGMENTS
We thank Juan Huerta for his technical
support in the management and maintenance of
the organisms in culture.
REFERENCES
Anson, M. L. (1938). The estimation of pepsin, tryp-
sin, papain and cathepsin with hemoglobin. Journal
of General Physiology, 22(1), 79–89. https://doi.
org/10.1085/jgp.22.1.79
Balon, E. K. (1986). Types of feeding in the ontogeny of
fishes and the life-history model. Environmental
Biology of Fishes, 16, 11–24. https://doi.org/10.1007/
BF00005156
Balon, E. K. (1990). Epigenesis of an epigeneticist: the
development of some alternative concepts on the
early ontogeny and evolution of fishes. Guelph Ichth-
yology Reviews, 1, 1–42.
Bradford, M. M. (1976). A rapid and sensitive method
for the quantization of microgram quantities of pro-
tein utilizing the principle of protein-dye binding.
Analytical Biochemistry, 72, 248–254. https://doi.
org/10.1016/0003-2697(76)90527-3
Chen, J. Y., Zeng, C., Jerry, D. R., & Cobcroft, J. M.
(2019). Recent advances of marine ornamental fish
larviculture: broodstock reproduction, live prey and
feeding regimes, and comparison between demersal
and pelagic spawners. Reviews in Aquaculture, 12(3),
1518–1541. https://doi.org/10.1111/raq.12394
Davis, B. J. (1964). Disc electrophoresis – II Method and
application to human serum proteins. Annals of the
New York Academy of Sciences, 121(2), 404–427.
https://doi.org/10.1111/j.1749-6632.1964.tb14213.x
Díaz-López, M., Moyano, F. J., Alarcón, F. J., García-Carre-
ño, F. L., & Navarrete del Toro, M. A. (1998). Cha-
racterization of fish acid proteases by substrate-gel
electrophoresis. Comparative Biochemistry and Phy-
siology B, 121(4), 369–377. https://doi.org/10.1016/
S0305-0491(98)10123-2
Del-Mar, E. G, Largman, C., Brodrick, J., & Geokas, M.
(1979). A sensitive new substrate for chymotrypsin.
Analytical Biochemistry, 99(2), 316–320. https://doi.
org/10.1016/S0003-2697(79)80013-5
Delbare, D., Lavens, P., & Sorgeloos, P. (1995). Clownfish
as a reference model for nutritional experiments and
determination of egg/larval quality. In P. Lavens
(Ed.), Fish and Shellfish Larviculture Symposium
(Vol. 24, pp. 22–25). European Aquaculture Society,
Special Publication.
Erlanger, B., Kokowsky, N., & Cohen, W. (1961). The
preparation and properties of two new chromo-
genic substrates of trypsin. Archives of Bioche-
mistry and Biophysics, 95(2), 271–278. https://doi.
org/10.1016/0003-9861(61)90145-x
García-Carreño, F. L., Dimes, L. E., & Haard, N. F. (1993).
Substrate-gel electrophoresis for composition and
molecular weight of proteinases or proteinaceous pro-
teinase inhibitors. Analytical Biochemistry, 214(1),
65–69. https://doi.org/10.1006/abio.1993.1457
Green, B. S., & McCormick, M. I. (2001). Ontogeny
of the digestive and feeding systems in the ane-
mone fish Amphiprion melanopus. Environ-
mental Biology of Fishes, 61, 73–83. https://doi.
org/10.1023/A:1011044919990
Gordon, A., Kaiser, H., Britz, P., & Hecht, T. (2000). Effect
of feed type and age-at-weaning on growth and
survival of clownfish Amphiprion percula (Poma-
centridae). Aquarium Science and Conservation, 2,
215–226. https://doi.org/10.1023/A:1009652021170
Gordon, A. K., & Hetch, T. (2002). Histological studies on
the development of digestive system of the clownfish
Amphiprion percula and the time of weaning. Journal
of Applied Ichthyology, 18(2), 113–117. https://doi.
org/10.1046/j.1439-0426.2002.00321.x
Khoa, T. N. D., Waqalevu, V., Honda, A., Shiozaki, K., &
Kotani, T. (2019). Early ontogenetic development,
digestive enzymatic activity and gene expression in red
sea bream (Pagrus major). Aquaculture, 512, 734283.
https://doi.org/10.1016/j.aquaculture.2019.734283
Khoa, T. N. D., Waqalevu, V., Honda, A., Shiozaki, K., &
Kotani, T. (2021). An integrative description of the
digestive system morphology and function of Japa-
nese flounder (Paralichthys olivaceus) during early
ontogenetic development. Aquaculture, 531, 735855.
https://doi.org/10.1016/j.aquaculture.2020.735855
11
Revista de Biología Tropical, ISSN: 2215-2075, Vol. 71: e51085, enero-diciembre 2023 (Publicado Mar. 02, 2023)
Khoo, M. L., Das, S. K., & Ghaffar, M. A. (2018). Growth
pattern, diet and reproductive biology of the clown-
fish Amphiprion ocellaris in waters of Pulau Tioman.
The Egyptian Journal of Aquatic Research, 44(3),
233–239. https://doi.org/10.1016/j.ejar.2018.07.003
Khoo, M. L., Das, S. K., & Ghaffar, M. A. (2019). Gastric
emptying and the enzymatic activity in the stomach
of Amphiprion ocellaris fed on artificial diet. Sains
Malaysia, 48(1), 1–6. http://dx.doi.org/10.17576/
jsm-2019-4801-01
Kolkovski, S. (2001). Digestive enzymes in fish larvae and
juveniles-implications and applications to formu-
lated diets. Aquaculture, 200, 181–201. https://doi.
org/10.1016/S0044-8486(01)00700-1
Laemmli, U. K. (1970). Cleavage of structural pro-
teins during the assembly of the head of bacte-
riophage T4. Nature, 227, 680–685. http://doi.
org/10.1038/227680a0
Lipscomb, T. N., Yanong, R. P., Ramee, S. W., & DiMa-
ggio, M. A. (2020). Histological, histochemical and
biochemical characterization of larval digestive
system ontogeny in black tetra Gymnocorymbus
ternetzi to inform aquaculture weaning protocols.
Aquaculture, 520, 734957. https://doi.org/10.1016/j.
aquaculture.2020.734957
Ma, Z., Guo, H., Zheng, P., Wang, L., Jiang, S., Qin, J. G.,
& Zhang, D. (2014). Ontogenetic development of
digestive functionality in golden pompano Trachino-
tus ovatus (Linnaeus 1758). Fish Physiology and Bio-
chemistry, 40(4), 1157–1167. https://doi.org/10.1007/
s10695-014-9912-0
Madhu, R., Madhu, K., & Retheesh, T. (2012). Life history
pathways in false clown Amphiprion ocellaris Cuvier,
1830: A journey from egg to adult under captive con-
dition. The Marine Biological Association of India,
54(1), 77–90.
Maroux, S., Louvard, D., & Baratti, J. (1973). The amino-
peptidase from hog-intestinal brush border. Biochimi-
ca et Biophysica Acta, 321(1), 282–295. https://doi.
org/10.1016/0005-2744(73)90083-1
Nazemroaya, S., Yazdanparast, R., Nematollahi, M. A.,
Farahmand, H., & Mirzadeh, Q. (2015). Ontogene-
tic development of digestive enzymes in Sobaity
sea bream Sparidentex hasta larvae under culture
condition. Aquaculture, 448, 545–551. https://doi.
org/10.1016/j.aquaculture.2015.06.038
Olivotto, I., Planas, M., Simoes, N., Holt, G. J., Avella, M.
A., & Calado, R. (2011). Advances in breeding and
rearing marine ornamentals. Journal of the World
Aquaculture Society, 42(2), 135–166. https://doi.
org/10.1111/j.1749-7345.2011.00453.x
Önal, U., Langdon, C., & Çelik, İ. (2008). Ontogeny of the
digestive tract of larval percula clownfish, Amphiprion
percula (Lacépède 1802): A histological perspective.
Aquaculture Research, 39(10), 1077–1086. https://
doi.org/10.1111/j.1365-2109.2008.01968.x
Peñaz, M. (2001). A general framework of fish ontogeny: A
review of the ongoing debate. Folia Zoologica-Praha,
50(4), 241–256.
Putra, D. F., Abol-Munafi, A. B., Muchlisin, Z. A., & Chen,
J. C. (2012). Preliminary studies on morphology and
digestive tract development of tomato clownfish,
Amphiprion frenatus under captive condition. Aqua-
culture, Aquarium, Conservation & Legislation, 5(1),
29–35.
Rodríguez-Ibarra, L. E., Abdo-de la Parra, M. I., Velas-
co-Blanco, G., & Aguilar-Zárate, G. (2017).
Desarrollo osteológico de la columna vertebral
y del complejo caudal de larvas del pez paya-
so Amphiprion ocellaris (Pomacentridae) en con-
diciones de cultivo. Revista de Biología Marina
y Oceanografía, 52(1), 113–119. http://dx.doi.
org/10.4067/S0718-19572017000100009
Rohini, K. M. V., Anil, M. K., & Neethu, R. P. (2018).
Larval development and growth of red saddleback
Anemonefish, Amphiprion ephippium, (Bloch, 1790)
under captive conditions. Indian Journal of Geo-
Marine Sciences, 47(12), 2421–2428.
Rønnestad, I., Yúfera, M., Ueberschär, B., Ribeiro, L.,
Sæle, Ø., & Boglione C. (2013). Feeding behaviour
and digestive physiology in larval fish: Current
knowledge, and gaps and bottlenecks in research.
Reviews in Aquaculture, 5, S59–S98. https://doi.
org/10.1111/raq.12010
Roux, N., Logeux, V., Trouillard, N., Pillot, R., Magré,
K., Salis, P., Lecchini, D., Besseau, L., Laudet, V.,
& Romans, P. (2021). A star is born again: Methods
for larval rearing of an emerging model organism,
the false clownfish Amphiprion ocellaris. Journal of
Experimental Zoology B, 336(4), 376–385. https://
doi.org/10.1002/jez.b.23028
Salze, G., McLean, E., & Craig, S. R. (2012). Pepsin onto-
geny and stomach development in larval cobia. Aqua-
culture, 324–325, 315–318. https://doi.org/10.1016/j.
aquaculture.2011.09.043
Shin, M. G., Ryu, Y., Choi, Y. H., & Kim, S. K. (2022).
Ontogenetic digestive physiology and expression
of nutrient transporters in Anguilla japonica lar-
vae. Aquaculture Reports, 25, 101218. https://doi.
org/10.1016/j.aqrep.2022.101218
Systat Software Inc. (2009). SigmaPlot (version 11.0).
Systa Software Inc., San Jose California, USA.
Versaw, W. S., Cuppett, D., Winters, D. D., & Win-
ters, L. E. (1989). An improved colorimetric assay
for bacterial lipase in nonfat dry milk. Journal
of Food Science, 54(6), 1557–1558. https://doi.
org/10.1111/j.1365-2621.1989.tb05159.x
12 Revista de Biología Tropical, ISSN: 2215-2075 Vol. 71: e51085, enero-diciembre 2023 (Publicado Mar. 02, 2023)
Walter, H. E. (1984). Proteinases: Methods with hemoglo-
bin, casein and azocoll as substrates. In H. U. Berg-
meyern (Ed.), Methods of enzymatic analysis (pp.
270–277). Chemic Weinham.
Weber, K., & Osborn, M. (1969). The reliability of
molecular weight determinations by dodecyl sulfate
polyacrylamide gel electrophoresis. Journal of Bio-
logical Chemistry, 244(16), 4406–4412. https://doi.
org/10.1016/S0021-9258(18)94333-4
Wilson, J. M., & Castro, L. F. C. (2010). 1–Morphologi-
cal diversity of the gastrointestinal tract in fishes.
Fish Physiology, 30, 1–55. https://doi.org/10.1016/
S1546-5098(10)03001-3
Yúfera, M., Moyano, F. J., & Martínez-Rodríguez, G.
(2018). The digestive function in developing fish
larvae and fry. From molecular gene expression to
enzymatic activity. In M. Yúfera (Ed.), Emerging
Issues in Fish Larvae Research (pp. 51–86). https://
doi.org/10.1007/978-3-319-73244-2_3
Yasir, I., & Qin, J. G. (2007). Embryology and early
ontogeny of an anemonefish Amphiprion ocellaris.
Journal of the Marine Biological Association of
the United Kingdom, 87(4), 1025–1033. https://doi.
org/10.1017/S0025315407054227
Zambonino-Infante, J. L., & Cahu, C. L. (2001). Onto-
geny of the gastrointestinal tract of marine fish
larvae. Comparative Biochemistry and Physio-
logy C, 130(4), 477–487. https://doi.org/10.1016/
S1532-0456(01)00274-5
Zambonino-Infante, J. L., & Cahu, C. L. (2007). Dietary
modulation of some digestive enzymes and metabolic
processes in developing marine fish: Applications to
diet formulation. Aquaculture, 268, 98–105. https://
doi.org/10.1016/j.aquaculture.2007.04.032
