19
Indian J. Fish., 62(2): 19-28, 2015
Captive breeding and developmental biology of Sahyadria denisonii
(Day 1865) (Cyprinidae), an endangered ish of the Western Ghats, India
T. V. ANNA MERCY, S. SAJAN AND V. MALIKA
Kerala University of Fisheries and Ocean Studies (KUFOS), Panangad, Kochi - 682 506, Kerala, India
e-mail: annamercy2012@gmail.com
ABSTRACT
Over the past few decades wild population of Sahyadria denisonii (Day 1865), an endemic ornamental barb of the Western
Ghats of India has been overexploited for aquarium trade and is presently listed under endangered (EN) category in the
IUCN Red List. The present study communicates the irst ever success of captive breeding and early developmental studies
of S. denisonii. Life history phases of S. denisonii were classiied into embryonic, larval, juvenile, subadult and adult stages.
Spawning season was from November to March in wild and fecundity varied depending on the size and age of breeding
pairs. Eggs were obtained through induced breeding using ovaprim hormone at 0.4 ml per kg body weight. Fertilised eggs
were adhesive, demersal and attached to any substratum having a diameter of 1184-1312 µm. Hatching took place 36 h after
fertilisation at a water temperature of 27.5±0.5ºC. At hatching, mean larval length was 3.5±0.2 mm with high amount of yolk
and the yolk sac remained up to 3-4 days. Organogenesis of larvae was completed 15-20 days after hatching. In this paper
full developmental sequence from egg to adult stages of S. denisonii in controlled condition is described.
Keywords: Captive breeding, Embryonic development, Miss Kerala, Sahyadria denisonii, Western Ghats
Introduction
The Western Ghats of India is one of the biodiversity
hotspots of the world (Myers, 2000) and its range of
hills running along India’s west coast (08º 19’08’’ - 21º
16’24’’N to 72º 56’24’’- 78º 19’40’’E) is one of the
richest regions in terms of its biological diversity. The
Western Ghats extends 1490 km from north to south with
a minimum width of 48 km and maximum of 210 km,
covering a total area of 136,800 km2 (Molur et al., 2011).
The area is drained by 38 east lowing and 27 west lowing
major rivers with running water and lacustrine habitats
(Abell et al., 2008). Ragahvan and Dahanukar (2013),
listed 320 species of freshwater ishes belonging to 11
orders, 35 families and 112 genera including certain
secondary freshwater species, which can also live in
brackishwater and marine habitats. The Kerala State
on the south - western corner of the Indian peninsula
is crisscrossed by 44 rivers arising from the Western
Ghats (41 west lowing and 3 east lowing), having an
immensely rich ichthyofauna of well over 300 species, of
which about 50% have ornamental and recreational value
(Mercy, 2009).
With the high demand and pricing of many ornamental
ish, they are being harvested at greater volumes and sold
at higher rates, threatening the viability and sustainability
of the resources (Chao, 2001; Vagelli and Erdmann, 2002;
Cato and Brown, 2003; Lunn and Moreau, 2004). In India,
most wild caught aquarium ish are from the Eastern
Himalayas and Western Ghats which are ecological
hotspots, known for their amazing freshwater biodiversity
and endemism (Allen et al., 2010; Molur et al., 2011).
Raghavan et al. (2013) reported that around two dozen
ishes are regularly being exported from Western Ghats
of Kerala, while Liya and Ramachandran (2013) listed
Tetraodon travancoricus, Dario dario, Sahyadria
denisonii, Botia striata and Carinotetraodon imitator as
the major species of ornamental ishes traded from India for
the years 2005-2010. Of India’s total live ornamental ish
exports to the tune of US $ 1.54 million during 2007-08,
S. denisonii accounted for almost 60-65%. Raghavan et al.
(2013) recorded that 310,791 numbers of S. denisonii
were exported from India during 2005-2012 and the main
markets were Singapore (48.63%), Hong Kong (30.52%)
and Malaysia (18.4%), with negligible quantities being
exported to Germany, United Kingdom and Japan.
Among the native ornamental ishes of Western Ghats
region, no species has received global fame and hobbyist
attention as that of the Redline torpedo ish, S. denisonii.
The distribution of S. denisonii is restricted to the southern
regions (Kerala and south Karnataka) of the Western
Ghats hotspot of India and their population appears
to be extremely fragmented in fourteen rivers namely
Chalakudy, Periyar, Achencovil, Pampa, Valapattanam,
Chaliyar, Kallar, Chandragiri, Bharathapuzha, Bhavani,
Karingode, Kuppam, Anjarkandipusha, Kuttiyadi
20
T. V. Anna Mercy et al.
(Biju et al., 2000; Shaji and Easa, 2001; Shaji et al., 2000;
Raghavan et al., 2010; Mercy et al., 2010 b; 2013 b).
Raghavan et al. (2010) and Silas et al. (2011) observed
that over-exploitation of freshwater ishes from the wild
for aquarium trade is the main reason for severe depletion
of their population. The existing export business of
freshwater ornamental ishes from India is not sustainable
since the present trade is completely dependent on wild
collection (Dahanukar et al., 2004; Raghavan et al., 2013).
Over the last few decades, wild population of S. denisonii
has declined due to various reasons (Dahanukar et al.,
2004; Mercy et al., 2010a; Raghavan et al., 2013) and
IUCN’s Freshwater Biodiversity Assessment of the
Western Ghats has categorised this species as Endangered
(Ali et al., 2010). In a bid to ensure long term availability
of S. denisonii, the government of Kerala is currently in the
process of inalising various measures aimed at checking
the uncontrolled, unorganised, unscientiic nature of wild
collection and export of this species (Raghavan et al.,
2010).
Captive breeding is a conservation strategy that
is widely used for the recovery and reintroduction
of endangered species (Kelley et al., 2006). Efforts
were taken to develop captive breeding technology for
indigenous ish species from Western Ghats of India
(Ogale, 2002; Padmakumar et al., 2004; Swain et al.,
2008; Mercy, 2009). It is envisaged that if S. denisonii
can be produced in captivity at a commercial level which
will deinitely boost the share of the species in the trade to
a large extent and will naturally lead to the conservation
of its germplasm. In this context, captive breeding
technology for S. denisonii was successfully developed
for the irst time and the early embryonic as well as larval
development stages were documented.
Materials and methods
Development of broodstock under simulated natural
condition
A water lowing habitat was constructed near the
Iruvanchipuzha, tributary of River Chaliyar, in such a
way that water from the same river is pumped to overhead
storage tank (10x10x2 m). Water was allowed to low
continuously through the artiicial habitat (10x3x2 m) by
gravitational force. All the materials used in the artiicial
habitat, such as sand, bottom sediments, rocks and plants
were brought from the same river from where the ishes
were collected. A diagrammatic representation of the
habitat is given in Fig. 1. Fifty juveniles of S. denisonii in
the size range 6-8 cm were collected from the Iruvanjipuzha
Tributary of River Chaliyar in Calicut District of southern
Kerala during September 2008. The collected ishes were
transported in oxygen illed polythene bags and stocked
in running water habitat. They were fed with protein rich
artiicial food, bloodworms and earthworms ad libitum.
Captive breeding
Twenty ive males and 15 females of S. denisonii
collected from the wild were segregated and reared in glass
tanks (3x2x2 ft.). As there is no distinct sexual dimorphism
in this species; the only dimorphic character observed
was that female has a slightly wider body than the male
when sexually mature. In a fully ripe female (size 10 cm
and above), eggs are seen extruded under slight pressure
and male ish (size 9 cm and above) is identiied by the
low of milt on applying gentle pressure on the abdominal
region towards the genital openings (Mercy et al., 2013b).
Maturity of the ripe eggs was determined by observing
the position of migratory nucleus of the eggs under a
microscope (Rottmann et al., 1991).
Induced breeding experiments were conducted
during December 2009 to March 2010, coinciding with
the natural spawning of this species in nature. From the life
history parameters, it was evident that the ish bred during
November-April months (Mercy et al., 2010a, 2013a,
Solomon et al., 2011). Since the broodishes are very
sensitive and handling stress can lead to their mortality,
the ish were anaesthetised before handling using clove
oil (30 mg l-1) (Sajan et al., 2012). The anaesthetised
male (10.4±1.6 cm, 15.5±2.7 g) and female (12.1±1.5 cm,
21.5±3.4 g) broodishes were injected intramuscularly
between the anterior region of dorsal in and lateral
line using a 1ml syringe with single dose of ovaprim
(SGnRH+ Domperidone- Syndel laboratory, Canada)
at a rate of 0.4 ml kg-1 body weight by late evening
(around 18.00 hrs). Breeding set comprising male and
female in 2:1 ratio were injected and were kept together
in glass tanks (150 l) provided with continuous aeration.
It was observed that induced broodishes did not release
eggs or milt by their own. After 10-12 h of latency period,
ish were inspected for readiness for ovulation. Both
males and females were stripped, and the eggs were dry
fertilised using milt collected (Harvey and Carolsfeld,
1993). Fertilised eggs were transferred to a glass tank
(10 l) and aerated gently. Ten trials were made successfully
during the season. Water quality parameters were
monitored at weekly intervals (Table 1). Embryonic and
larval developments were documented and photographed
with binocular stereo microscope (LABOMED) with
digital camera (Canon Power Shot-A570).
The hatchery produced juveniles (F1 generation)
were reared in the hatchery at Thiruvambady and hundred
numbers of juveniles were transported live to the hatchery
at the College of Fisheries, Ernakulam during June 2010.
They were acclimatised and reared in outdoor rectangular
cement tanks (10 t capacity) and were fed with the same
food as before. Water quality parameters were maintained
similar to the wild (Table 1). During January 2012, three
21
Kerala
River & Lakes
N
Arabian
Sea
Inlet
Outlet
Fig. 1. Schematic representation of the modiication of habitat for captive breeding of Sahyadria denisonii
Table 1. Water quality parameters of the wild habitat and broodstock
rearing tank
Parameters
Range
Water temperature (oC)
26 - 28
pH
7.0 -7.5
Dissolved oxygen (mg l-1)
5.0- 6.8
Total alkalinity (mg l-1)
20 - 25
Hardness (mg l-1)
20 - 25
Ammonia (mg l-1)
<0.01
Nitrate (mg l-1)
<0.01
pairs of S. denisonii of F1 generation were successfully
bred following the same protocol as above. Instead of
clove oil, MS-222 (150 mg l -1 ) was used as anaesthetic
agent (Mercy et al., 2013a)
Results and discussion
The results of this study gave a better understanding
about the breeding protocol of S. denisonii. In hatchery,
broodstock management is one of the major aspects for
successful induced breeding of any ish species. Proper
care of broodstock is very important for assuring the
production of eggs, fry and ingerlings (Siddik et al.,
2013). In the present study, S. denisonii was successfully
bred under captivity by artiicial fertilisation, using
ovaprim as inducing agent. Ovaprim was used as
an inducing agent for Labeo dussumieri (Kurup,
1994); Channa striatus (Haniffa et al., 2000); Systomus
22
Captive breeding and developmental biology of Sahyadria denisonii
sarana (Chakraborty et al., 2003); Mystus montanus
(Arockiaraj et al., 2003), Schizothorax richardsonii
(Agarwal et al., 2007); Garra surendranathanii (Thamby,
2009) and Dwakinsia ilamentosus (Mercy, 2009).
Ovaprim has been accepted as the most superior inducing
agent for carps with high spawning success, percentage
of fertilisation and hatching rate (Nandeesha et al., 1990).
Fertilised eggs (Fig. 2.2) of S. denisonii were heavily
yolked, transparent, spherical and yellow in colour. The
eggs were sticky and seen stuck to the glass surface
similar to Puntius species. Egg incubation and hatching
is better performed in glass tanks under slight continuous
aeration. According to Balon (1975), S. denisonii is a
lithophil (morphotype), open substratum spawner and
falls under ethological class of non-guarders as per the
eco-morphological classiication. According to Mercy et al.
(2013b) absolute fecundity of S. denisonii ranged from
293 to 967ova per female, while Solomon et al. (2011)
recorded it as 376 to 1098.
The embryonic period started just after fertilisation
and ended when the embryo acquired all the organ systems
as in other ishes (Table 2; Fig. 2). In the present breeding
trial, latency period was observed as 12.78±0.83 h,
fertilisation rate as 86.11±5.23% and hatching rate as
85.89±2.98%. The higher rates of artiicial fertilisation
during the present investigation may be related to the
dry method of fertilisation where the viability of the
spermatozoa remains high. However, the percentage of
fertilisation is also related to the maturity and weight of
ish (Haque and Ahmed, 1991; Agarwal et al., 2007). In
the present study, breeding was performed at an ambient
temperature of 26.0-28.0°C. This range of temperature
is suitable for breeding of most indigenous small ishes
(Islam and Chowdhury, 1976; Akhteruzzaman et al., 1992;
Siddik et al., 2013).
Embryonic phases
Formation of embryo : The fertilised eggs (Fig. 2.2) were
spherical, orange brown in colour and adhesive with a
size of 1184-1312 µm dia. Fertilised eggs were sticky
due to the sticky secondary membrane from the follicular
envelope. Egg development started immediately after
fertilisation and it activated the cytoplasmic movements.
Yolk free cytoplasm begins to stream towards the animal
pole gradually segregating the blastodisc from the vegetal
cytoplasm. Within 20 min, the streaming movement
of the protoplasm towards the lower pole is completed
and a blastodisc is formed (Fig. 2.4). The irst cleavage
commenced 20 min after fertilisation when the blastodisc
was divided into two blastomeres (Fig. 2.5) followed
by second cleavage at the 40th min after fertilisation
(Fig. 2.6). The sixteen cell stage (Fig. 2.8) was noticed
within 80 min and 32 cell stage at 100-110 min
post-fertilisation. Cell division continued rapidly after the
eight blastomere stage (Fig. 2.7). Morula (Fig. 2.9) and
blastula (Fig. 2.10) stage occurred at 180th and 210th min
respectively.
Cell division took place more or less synchronously
during the early stages of the blastula period. Blastoderm
Table 2. Embryonic developmental stages of Sahyadria denisonii
Developmental stages
Duration
Figure
Features
Eggs freely ooze out
Fertilized eggs
00:00 h.
00:00 h
Sticky and orange in colour
Spherical, orange coloured, demersal and adhesive
Blastodisc stage
Two cell stage
Four cell stage
Eight cell stage
Sixteen cell stage
Morula stage
Blastula stage
Dome stage
Late blastula stage
Blastopore stage
Yolk plug stage
00:10 h
00:20 h
00:40 h
00:60 h
01:20 h
03:00 h
03:30 h
06:00 h
08:00 h
08:30 h
09:00 h
2.01
2.02
2.03
2.04
2.05
2.06
2.07
2.08
2.09
2.10
2.11
2.12
2.13
2.14
Head - tail bud stage
Tail free stage
Eye vesicle stage
Twitching stage
Just before hatching
Hatching
17:00 h
18:00 h
19:00 h
30:00 h
34:00 h
36:00 h
Just hatched larvae
Hatchling
3.54 ± 0.21 mm TL
36:00 h
36:00 h.
2.16
2.17
2.18
2.19
2.20
5.21
2.22
2.23
2.24
2.25
Cytoplasm streams toward animal pole
First cleavage
Second cleavage
Third cleavage
Fourth cleavage
Blastodermal cells formed after cleavage, arranged in group in the animal pole
Crown like blastodern spread over yolk at animal pole
Bblastoderm expanded to the equator of the yolk sphere; pre-gastrula stage
Blastoderm extended towards the vegetal pole
Large yolk plug protrudes from the blastopore.
Yolk invasion completed by gradual spreading over the germ layer. Rudimentary head and tail
appeared and became differentiated.
Head and tail rudiment visible
Yolk at the tail end elevated to inside to free tail from yolk mass.
Eye vesicle observed
Vigorous movements before hatching.
Continuous beating of the caudal region especially around middle region of the body
The egg shell breaks up and the tail emerges out irst, followed by the head; 3.54 ± 0.21 mm in total
length
Larvae hatch out from egg capsule
Newly hatched larvae non-pigmented and had an average total length of 3.54 ± 0.21 mm and yolk sac
of 1 mm dia.
23
T. V. Anna Mercy et al.
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
Fig. 2. Embryonic and larval development stages of Sahyadria denisonii
38
39
Captive breeding and developmental biology of Sahyadria denisonii
was observed at 5 h post-fertilisation, (hpf) followed
by dome stage (Fig. 2.11) at 1 h. At dome stage, the
blastoderm formed a dome-like shape due to bulging
of the yolk towards the animal pole and the epiboly
continued during the gastrulation period. In the gastrula
period, extensive cell movements were observed,
including involution, convergence and extension,
producing the three primary germ layers and the
embryonic axis. Gastrulation began with cell involution
at around 50% epiboly. At 75% epiboly stage (Fig. 2.12),
the embryonic shield became less distinctive, as compared
to shield stage and its cells got repacked to elongate
the shield along the animal pole axis. At 90% epiboly
stage, the yolk plug was clearly seen in the vegetal pole
(Fig. 2.13 and 2.14) and was noticed at 9 hpf. The
segmentation period was observed at 15 h which was
characterised by the sequential formation of the somites,
and this period lasted up to hatching. During this period,
the embryo elongated along the animal pole axis, the tail
bud became longer and rudiments of the primary organs
became visible (Fig. 2.15).
Differentiation of embryo: The head and tail bud of the
embryo (Fig. 2.16) became clearly distinguishable after
17 h. In this stage, the optic primordium had a prominent
horizontal crease and tail bud became more prominent.
At 19 hpf, the eye vesicle was observed, the embryo
increased in size and eighteen myotomes were clearly
visible (Fig. 2.17). Tail bud started to separate from the
yolk sac at 22 hpf (Fig. 2.18). The yolk extension got
clearly delimited from the yolk ball as the tail straightened
out. Twitching movements of the embryo started by 23
hpf. At 26 h, tail end was free from the rest of the yolk
sac (Fig. 2.19) and at 30 h, embryo started to twist or
rotate completely inside the egg membrane. After 34
h, movements of the embryo within the egg membrane
were very frequent. Close to hatching, the twitching and
lashing of the embryo inside the egg capsule became
rapid (Fig. 2.20). The egg shell broke at 36 h due to rapid
shaking movements of the body and the head emerged
out irst, followed by the tail (Fig. 2.21 to 2.24). Hatching
of eggs continued up to 40 h, probably the temperature
and oxygenation of the water played an important role
in incubation as reported by Laurila et al. (1987) and
Dwivedi and Zaidi (1983).
Larval phase
Larval phase can be further subdivided into yolk sac
stage, pre-lexion stage, lexion stage and post-lexion
stage (Fig. 2.25 to 2.39) according to the classiication by
Pena and Dumas (2009).
Yolk sac larva/eleuthero embryo: The newly hatched
larvae (Fig. 2.25) were non-pigmented with an average
total length of 3.5±0.2 mm and yolk sac of 1mm dia.
24
Hatchlings had non- pigmented eyes and were devoid of
distinct mouth and ins. Head seemed to be in contact with
yolk sac and optic vesicles. The yolk sac is large and was
found attached to the body along its ventral side. A narrow
in fold was found to be differentiated as a thin membrane,
surrounding the caudal region and extended up to the
yolk sac. The crawling movements of the hatchlings were
detected immediately after hatching.
Two day old larva: On the second day, the larvae appeared
to be more active; and inhabited the bottom of the tank as
a group with average total length of about 4.1±0.2 mm
(Fig. 2.28). The head portion seemed to be further
developed. On closer observation, the heart was seen
pulsating rhythmically. Pigmentation started in the eyes
(Fig. 2.26). Normally in ish, the shape of the yolk sac
undergoes signiicant changes during organogenesis
(Winnicki et al., 2001). In case of S. denisonii during
embryogenesis, the yolk sac got divided into two parts,
proximal portion with spherical shape and caudal part
with cylindrical shape. Yolk sac became slightly reduced
in size, with 3 mm in length towards the ventral side of
the body (Fig. 2.27). Ventral and caudal in folds were
separated by the formation of anus. Formation of pectoral
in folds was also seen.
Three day old larva: Larvae on the third day, reached
an average length of 4.5±0.2 mm. Yolk sac was seen
further reduced. Mouth and anus became distinguishable.
Pectoral ins with in rays started to move vigorously
(Fig. 2.29). Head became more prominent and free
movements of the eye balls were also noticed
Melanophores appeared around the head and snout
(Fig. 2.30). Larvae exhibited vigorous movements
upwards to the column water and inally sank to the
bottom. All the active larvae got accumulated together into
a lump-like ball showing strong thigmotactic behaviour.
Pre-lexion Stage : The pre-lexion stage began on day 4
when the larva started its irst external feeding. Average
length (TL) at this stage was 5.1±0.2 mm. Pre-lexion stage
extended from a period of about 4 to 8 days; till it reached
an average TL of 6.8±0.4 mm. Yolk sac got reduced and
became tube like with 2.2 mm length and 0.5 mm width.
At the onset of the pre-lexion stage the eye became fully
pigmented and the mouth and anus opened. Alimentary
canal was found to contain a yellowish luid. Air bladder
was noticed and occurrence of pigmentation on the air bladder
started (Fig. 2.31 to 2.32). Opercular movements was also
noticed. Caudal in became modiied with rudimentary
in rays. Pigments were found more concentrated on the
anterior region of the body and also in caudal in fold.
Vigorous movements of the mouth was noticed as if the
larva was in search of food. Eye balls exhibited rotating
movements and larvae showed grouping behaviour when
25
T. V. Anna Mercy et al.
all of them accumulated into a bundle towards the latter
stage. Melanophores also appeared in the middle and in
the notochord region. One week old larva was very active
and started free swimming. Egg yolk was completely
absorbed at this time. Average body length of the larvae
increased to 7 mm and at this stage larvae fed entirely on
exogenous food.
Flexion stage: The lexion stage started from day 08
(average TL 07.4±0.3 mm) and extended to day 17
(average TL 10.4±1.5 mm). The complement of the
notochord lexion was characterised by the anterioposterior orientation of the caudal rays. A key event
was the development of lexion in the ventral side of
the spinal cord in the notochord associated with the tail
in. On day 9 or 10, the single chambered air bladder
(Fig. 2.32) was transformed into double chambered
(Fig. 2.33). Furthermore, together with the disappearance
of the larval in fold and development of the ins a change
in swimming style was also observed (Fig. 2.34). Similar
observations were also reported in other teleost larvae
(Van Snik et al., 1997; Gisbert, 1999; Gisbert et al., 2002).
Post-lexion stage: The post-lexion stage was observed
from day 17 (average TL 10.4±1.5 mm) and extended to
day 26 (average TL 16.7±2.1 mm). After two weeks, the
larvae completely metamorphosed; and the average body
length increased to 11.2±1.5 mm. Changes during this
stage included the development of dorsal in having 6 - 7
in rays. A vertical black band was formed over the body
and an inferior mouth was clearly observed. Fin elements
became evident in the dorsal, caudal, anal and pelvic
ins during the post-lexion stage. Body pigmentation
increased in the middle part of the body.
Larval feeding and weaning : The newly hatched larvae
(3.5±0.2 mm) fed on its yolk content up to 3 to 4 days after
hatching. Infusoria (pure culture of Paramecium) was
given as irst exogenous feed. The larvae were fed with
freshly harvested paramecium at every 3 h interval. About
40-50% water exchange (30 l glass tanks) was done every
day after exogenous feeding. After 10 days, the larvae were
stocked in glass tanks (50 l) with an approximate stocking
density of 10 larvae l-1 and were fed with live micro-worms
(Panagrellus spp.) cultured using bread or oats medium.
No substratum was provided during irst two weeks of
larval rearing and thereafter larval tanks were provided
with 2 or 3 small pieces of clean rock as substratum for
the larvae. Hatchlings were gradually weaned to artiicial
feeds (Artemia lakes), 12-14 days after hatching. Uneaten
food was siphoned out from the bottom of each tank every
morning before irst feeding and dead ish were counted
and preserved to analyse percentage of mortality.
Dissolved ammonia plays a vital role during larval
rearing of S. denisonii and daily water quality assessment
is necessary. In nature, post-larval stages of S. denisonii
exhibit a bottom dwelling habit and they rely on bottom
deposited diets such as detritus, algae and diatom that
grows on pebbles and stones. Similar indings were
also noticed by Costa and Fernando (1967). Feeding
experiments suggested that artiicial feed can be used
for rearing post-larvae as well as juvenile stages of
S. denisonii in aquaria (Mercy and Sajan, 2014) and it will
be beneicial for the development of better larval rearing
techniques for S. denisonii under hatchery condition.
Juvenile phase
One month old juveniles (average TL 29.8±2.3 mm) of
S. denisonii showed vertical body banding pattern.
Juveniles of S. denisonii possessed four vertical black
cross band at nuchal, sub dorsal, supra anal and caudal
positions (Fig. 2.35 to 2.36). Later, after disappearance of
vertical cross bands, a lateral black bar started to appear
from posterior margin of the eye region and extended
towards the caudal peduncle. A transverse black and
yellow band was observed in the tip of each lobe of the
caudal in. The anterior base of dorsal in exhibited a black
and red blotch (Fig. 2.36).
Sub-adult and adult phase
The juveniles (Fig. 2.37) reached an average TL of
35.8±5.3 mm size after rearing for three to four months
in iber tanks (200x100x50 cm). A red streak started to
appear from the tip of the snout, parallel to the black
stripe already formed posteriorly. It extended up to the
middle of body below the dorsal in. This sub-adult stage
(Fig. 2.38) externally resembled adults in presence of
these stripes. The four vertical cross bands disappeared
and their body became greenish brown on the dorsal
side and silvery in the ventral side (Fig. 2.38). Juveniles
and sub-adults kept under ibre tank condition and in
artiicial habitat exhibited size variation during growth.
This may be due to the availability of natural food and
environmental conditions as has been reported by Hecht
and Pienaar (1993) and Wankowski and Thorpe (1979) in
Salmo salar. The ish became adult (Fig. 2.39) on reaching
a size of TL 7.8±2.2 cm.
The results clearly demonstrated the possibility
of using both clove oil and MS-222 as anaesthetics and
synthetic ish breeding hormone ovaprim for effective
induced breeding and seed production of S. denisonii
under captive condition. This captive breeding technology
has also been transferred to the ornamental ish breeders
from different states by the Marine Product Export
Development Authority, Kochi (Anon., 2014). The present
work generated information on the early life history and
developmental stages and also on commencement of irst
feeding time for larval rearing. The indings of the present
Captive breeding and developmental biology of Sahyadria denisonii
26
study can be used in induced breeding of S. denisonii in
hatcheries and in conservation of this critically endangered
valuable species.
Cato, J. C. and Brown, C. L. 2003. Marine ornamental
species: Collection, culture, and conservation. Iowa State
University Press, Ames, Iowa, 310 pp.
Acknowledgements
Chakraborty, B. K., Miah, M. I., Mirza, M. J. A. and Habib,
M. A. B. 2003. Rearing and nursing of local Sarpunti,
Puntius sarana at different stocking densities. Pakistan
J. Biol. Sci., 6 (9): 797-800.
We express our sincere thanks to Marine Products
Export Development Authority (MPEDA), Kochi for the
inancial assistance (2007-2010). We are also grateful
to the Dean, College of Fisheries, Kerala University of
Fisheires and Ocean Studies (KUFOS) for providing the
necessary facilities to carry out this work. Thanks are
due to anonymous reviewers for their criticisms which
improved the quality of the manuscript.
References
Chao, N. L. 2001. The ishery diversity and conservation
of ornamental ishes in the Rio Negro Basin, Brazil:
A review of Project Piaba (1989 - 1999). In: Chao, N. L.,
Petry, P., Prang, G., Sonnenschein L. and Tlusty M. T.
(Eds.), Conservation and management of ornamental
ish resources of the Rio Negro Basin, Amazonia Brasil
- Project Piaba. Editora da Universidade do Amazonas,
Manaus, p.161-204.
Abell, R., Allan, J. D. and Lehner, B. 2007. Unlocking the
potential of protected areas for freshwaters. Biol. Conser.,
134: 48-63.
Costa, H. H. and Fernando, E. C. M. 1967. The food and feeding
relationship of the common meso and macrofauna in the
Mahaoyam a small mountaimous stream at peracniya.
Ceyl. J. Sci., 7: 54-90.
Agarwal, N. K., Thapliyal, B. L. and Raghuvanshi, S. K. 2007.
Induced breeding and artiicial fertilisation of snow trout,
Schizothorax richardsonii through the application of
ovaprim. J. Inland Fish. Soc. India, 39(1): 12-19.
Dahanukar, N. and Raghavan, R. 2007. Freshwater ishes of
the Western Ghats: Checklist. FFSG Newsletter-Min.,
1(8): 6-16.
Akhteruzzaman, M., Kohinoor, A. H. M. and Shah, M. S. 1992.
Observations on the induced breeding of Puntius sarana
(Ham). Bangladesh J. Zool., 20: 291-295.
Ali, A., Raghavan, R. and Dahanukar. N. 2010. Puntius
denisonii In: IUCN 2011. IUCN Red List of threatened
species. Version 2011.1. www.iucnredlist.org (Accessed
19 August 2011).
Allen, D. J., Molur, S. and Daniel, B. A. 2010. The status and
distribution of freshwater biodiversity in the Eastern
Himalaya. IUCN, Cambridge, UK and Gland, Switzerland.
89pp,
Anon., 2014. Technology transfer by hands-on training on captive
breeding. http://isheriesexploration.blogspot.in/2014/08/
technology-transfer-by-hands-on.html (Accessed August
2014).
Arockiaraj, A. J., Haniffa, M. A., Seetharaman, S. and Singh, S. P.
2003. Early development of a threatened freshwater
catish Mystus montanus (Jerdon). Acta Zool. Taiwanica,
14(1): 23 -32.
Balon, E. K. 1975. Reproductive guilds of ishes - proposal and
deinition. J. Fish. Res. Board Canada, 32: 821-864.
Biju, C. R., Thomas, K. R. and Ajitkumar, C. R. 2000. Index
areas selected for long time monitoring and conservation.
In: Chhapgar, B. F. and Manakadan, R. (Eds.), Ecology of
hill streams of the Western Ghats with special reference
to ish community, inal report 1996-1999. Project report
submitted to Bombay Natural History Society, Bombay,
India, p. 34-48.
Dahanukar, N., Raut, R. and Bhat, A. 2004. Distribution,
endemism and threat status of freshwater ishes in the
Western Ghats of India. J. Biogeogr., 31(1): 123-136.
Dwivedi, S. N. and Zaidi, S. G. S. 1983. Development of carp
hatcheries in India. Fishing Chimes, 3(5): 29-37.
Gisbert, E. 1999. Early development and allometric growth
patterns in Siberian sturgeon and their ecological
signiicance. J. Fish Biol., 54: 852-862.
Gisbert, E., Merino, G., Muguet, J. B., Bush, D., Piedrahita, R. H.
and Conklin. D. E. 2002. Morphological development and
allometric growth patterns in hatchery-reared California
halibut larvae. J. Fish Biol., 61: 1217-1229.
Haniffa, M. A., Merlin Rose, T. and Francis, T. 2000. Induced
spawning of the striped Murrel Channa striatus using
pituitary extracts, human chorionic gonadotropin,
luteinising hormone releasing hormone analogue, and
ovaprim. Acta Icht. Piscat., 30(1): 53-60.
Haque, M. T. and Ahmed, A. T. A. 1991. Breeding biology
of tawes (Puntius gonionotus Bleeker). Indian J. Fish.,
38(1): 26-29.
Harvey, B. and Carolsfeld, J. 1993. Induced breeding in tropical
ish culture. International Development Research Centre,
Ottawa, 144 pp.
Hecht, T. and Pienaar, A. G. 1993. A review of cannibalism and
its implications in ish larviculture. J. World Aquacult. Soc.,
24(2): 246-261.
Islam, Q. and Chowdhury, A. Q. 1976. Induced spawning of
major carp for commercial production of fry in the ish
seed farm. Bangladesh J. Zool., 4: 51-61.
T. V. Anna Mercy et al.
Kelley, J. L., Magurran, A. E. and Macias Garcia, C. 2006.
Captive breeding promotes aggression in an endangered
Mexican ish. Biol. Conserv., 133: 169-177.
Kurup, B. M. 1994. Maturation and spawning of an indigenous
carp Labeo dussumieri (Va1.) in the river Pamba.
J. Aquacult. Trop., 9: 119-132.
Laurila, S., Piironen, J. and Holopainen, I. J. 1987. Notes on
egg development and larval and juvenile growth of crucian
carp Carassius auratus. Ann. Zool. Fennici., 24: 315- 321.
Liya, J. and Ramachandran, A. 2013. Major sustainability issues
and comparative sustainability assessment of wild caught
indigenous ornamental ishes exported from Kerala, India.
Fish. Technol., 50: 175-179.
Lunn, K. E. and Moreau, M. A. 2004. Unmonitored trade in
marine ornamental ishes: the case of Indonesia's Banggai
cardinalish, Pterapogon kauderni. Coral Reefs, 23: 344-351.
Myers, N., Mittermeier, R. A., Mittermeier, C. G., da Fonseca,
G. A. B. and Kent, J. 2000. Biodiversity hotspots for
conservation priorities, Nature, 403: 853 – 858.
Mercy, T. V. A. 2009. Status of standardised breeding and
propagation technology of indigenous ornamental ishes
of Western Ghats of India. J. World Aquacult. Soc., 35(4):
40-42.
Mercy, T. V. A., Malika, V. and Sajan, S. 2010a. Reproductive
biology and captive breeding of Puntius denisonii, an
indigenous ornamental ish of Western Ghats of India.
India International Ind-Aquaria, Marine Product Export
Development Authority, Chennai, Tamil Nadu, India.
Mercy, T. V. A., Malika, V. and Sajan, S. 2010b. Breakthrough
in breeding of Puntius denisonii. INFOFISH, 3(4): 13-17.
Mercy, T. V. A., Malika, V. and Sajan, S. 2013a. Use of tricaine
methanesulfonate (MS 222) to induce anaesthesia in
Puntius denisonii (Day, 1865) (Teleostei: Cypriniformes:
Cyprinidae) - an endangered barb of the Western Ghats
Hotspot, India. J. Threat. Taxa., 5(9): 4414-4419.
Mercy, T. V. A., Malika, V. and Sajan, S. 2013b. Reproductive
biology of Puntius denisonii (Day 1865) - an endemic
ornamental cyprinid of the Western Ghats of India. Indian
J. Fish., 60(2): 73-78.
Mercy, T. V. A. and Sajan, S. 2014. Early growth performance
of an endangered barb Sahyadria denisonii (Day, 1865)
fed with different diets under controlled conditions. Fish.
Technol., 51(4): 286-290.
Molur, S., Smith, K. G., Daniel, B. A. and Darwall, W. R. T.
2011. The status and distribution of freshwater biodiversity
in the Western Ghats. International Union for Conservation
of Nature (IUCN), Gland, Switzerland, 25 pp.
Nandeesha, M. C., Rao, K. G., Jayanna, R., Parker, N. C.,
Varghese, T. J., Keshavnath, P. and Shetty, H. P. C. 1990.
27
Induced spawning of Indian major carps through single
application of ovaprim. In: Hirani. R. and Hanyu, I. (Eds.),
Proceedings of the the Second Asian Fisheries Forum,
Asian Fisheries Society, Manila, Philippines, p. 581-585.
Ogale, S. N. 2002. Mahseer breeding and conservation, and
possibilities for commercial culture: the Indian experience.
In: Petr, T. and Swar, S. B. (Eds.), Coldwater isheries in
the Trans Himalayan Countries. FAO Fisheries Technical
Paper 431, p. 193-212.
Padmakumar, K. G., Krishnan, A., Bindu, L., Sreerekha, P. S.
and Joseph, N. 2004. Captive breeding for conservation
of endemic ishes of Western Ghats, India. National
Agriculture Technology Project (NATP), Kerala
Agricultural University, India, 79 pp.
Pena, R. and Dumas, S. 2009. Development and allometric growth
patterns during early larval stages of the spotted sand bass
Paralabraxmaculato fasciatus (Percoidei: Serranidae).
In: Clemmesen, C., Malzahn, A. M., Peck, M. A. and
Schnack, D. (Eds.), Advances in early life history study of
ish, Scientia Marina 73S, Barcelona (Spain), p. 183-189.
Raghavan, R., Dahanukar, N., Tlusty, M., Rhyne, A., Kumar, K. K.,
Molur, S. and Rosser, A. 2013. Uncovering an obscure
trade: threatened freshwater ishes and the aquarium
markets. Biol.Conserv.,164: 158-169.
Raghavan, R., Prasad, G., Ali, P. H., Pereira, B., Baby, F. and
Ram Prasanth, M. 2010. Miss Kerala added to the IUCN
Red List of Threatened Species. Current Sci., 98(2): 132.
Rottmann, R. W., Shireman, J. V. and Chapman, F. A. 1991.
Determining sexual maturity of broodstock for induced
spawning of ish. SRAC Publication No.423, 4 pp.
Sajan, S., Mercy, T. V. A. and Malika, V. 2012. Use of an
eco-friendly anaesthetic in the handling of Puntius
denisonii (Day, 1865) - an endemic ornamental barb of the
Western Ghats of India. Indian J. Fish., 59(3): 131-135.
Shaji, C. P. and Easa, P. S. 2001. Freshwater ishes of the Western
Ghats - A ield guide. Kerala Forest Research Institute
(KFRI), Peechi and National Bureau of Fish Genetic
Resources (NBFGR), Lucknow, India, 109 pp.
Shaji, C. P., Easa, P. S. and Gopalakrishnan, A. 2000. Freshwater
ish diversity of Western Ghats. In: Ponniah, A. G. and
Gopalakrishnan A. (Eds.), Endemic ish diversity of Western
Ghats, NBFGR-NATP publication 1, National Bureau of
Fish Genetic Resources, Lucknow, India, p. 33-35.
Siddik, M. A. B., Nahar, A. L., Ahamed, F., Masood, Z. and
Hossain, M. Y. 2013. Conservation of critically endangered
olive barb Puntius sarana (Hamilton, 1822) through
artiicial propagation. Our Nature, 11(2): 96-104.
Silas, E. G., Gopalakrishnan, A., Ramachandran, A., Anna
Mercy, T. V.., Kripan Sarkar., Pushpangadan, K. R., Anil
Kumar, P., Ram Mohan, M. K. and Anikuttan, K. K. 2011.
Guidelines for green certiication of freshwater ornamental
Captive breeding and developmental biology of Sahyadria denisonii
28
ish. Marine Product Export Development Authority,,
Kochi, Kerala, 105 pp.
1996). Ph. D. Thesis, Cochin University of Science and
Technology, Kerala, 177pp.
Solomon, S., Ramprasanth, M. R., Baby, F., Pereira, B., Tharian,
J., Ali, A. and Raghavan, R. 2011. Reproductive biology of
Puntius denisonii, an endemic and threatened aquarium ish
of the Western Ghats and its implications for conservation.
J. Threat. Taxa, 3(9): 2071-2077.
Vagelli, A. A. and Erdmann, M. V. 2002. First comprehensive
ecological survey of the Banggai cardinal ish, Pterapogon
kauderni. Environ. Biol. Fishes, 63: 1-8.
Swain, S. K., Chakrabarty, P. P. and Sarangi, N. 2008. Export and
breeding protocols developed for the indigenous ishes of
North-east India. In: Kurup, B. M., Boopendranath, M. R.,
Ravindran, K., Saira Banu and Nair, A. G. (Eds.), Book on
ornamental ish breeding, farming and trade. Department
of Fisheries, Govt. of Kerala, India, p. 114-134.
Thamby, S. 2009. Bionomics, cryopreservation of gametes
and captive breeding behaviour of threatened hill stream
cyprinid, Garra surendranathanii (Shaji, Arun & Easa,
Date of Receipt
:
20.04.2013
Date of Acceptance :
16.10.2014
Van Snik, G. M. J., Van den Boogaart, J. G. M. and Osse, J. W.
M. 1997. Larval growth patterns in Cyprinus carpio and
Clarias gariepinus with attention to the fanfold. J. Fish
Biol., 50: 1339-1352.
Wankowski, J. W. J. and Thorpe, J. E. 1979. The role of food
particle size in the growth of juvenile Atlantic salmon
(Salmo salar L). J. Fish Biol., 14: 351-370.
Winnicki, A., Korzelecka-Orkisz, A., Bonisławska, M. and
Formicki, K. 2001. Bipartity of the yolk sac in cyprinid
embryos. Arch. Ryb. Pol., 9(2): 279-286.