E3S Web of Conferences 147, 01005 (2020)
3rd ISMFR

https://doi.org/10.1051/e3sconf/202014701005

Dietary Value of Waste-Fed Rotifer Brachionus
rotundiformis on the Larval Rearing of JapaneseWhiting Sillago japonica
Erick Ochieng Ogello1, Stenly Wullur2*, Yoshitaka Sakakura3, and Atsushi Hagiwara3,4
1

Department of Fisheries and Natural Resources, Maseno University, P.O.Box Private Bag, Maseno,
Kenya
2Faculty of Fisheries and Marine Science, Sam Ratulangi University, Manado 95115, Indonesia
3
Graduate School of Fisheries and Environmental Sciences, Nagasaki University, 1-14 Bunkyo,
Nagasaki 852-8521, Japan
4
Institute of Integrated Science and Technology, Nagasaki University, Japan

Abstract. Live food resources are useful for larval fish rearing. However,production of sufficient live food resources is expensive. This study
employed a cost-effective rotifer culture technique using fish waste diet(FWD) and investigate the effect of the FWD-fed rotifer on larval rearing of
the Japanese whiting, Sillago japonica. Fertilized eggs of S. japonica were
hatched in polycarbonate tanks containing 100 l of artificial seawater at 10
eggs l-1 with 50 ml min-1 of aeration at room temperature. Two diets (i.e.
FWD-fed rotifers and rotifers fed with super fresh Chlorella-V12 as control)
were used with 10 rotifers ml-1 for 10 days. Fish were sampled every twodays for morphometric and gut content analysis. Fatty acid analysis wasdone for both rotifers and fish larvae. The fish larvae fed with FWD-rotifers
had higher total length than those given control diet. There was no
significant difference in survival rate, viability, dry weight, gut content, headlength, eye diameter, and body depth between the two diets. The DHA
recorded of total lipid for the fish given FWD-rotifer and control fish are3
5.2% and 18.2% respectively. The use of waste-fed rotifers is cost-effective
method to enhance the production of larval fish rearing in hatcheries.

1 Introduction
At the close of endogenous feeding phase, most marine larval fishes still have small mouth
gap and rudimentary digestive track and therefore, cannot ingest and digest inert feed [1, 2].
At this critical phase, the larval fishes require timely and adequate supply of appropriate live
feed that are readily ingested, efficiently digested and with essential nutrition for their healthy
growth and development [3]. Depending on the rotifer culture method and condition,
essential nutrients may be deficient, sometimes lower than the required threshold for healthy
fish development [4]. Therefore, due to the possibility of nutritional deficiencies, rotifers areoften provided with supplements to improve their nutritional value for larviculture. The
freshly cultured microalgae, sometimes enriched with essential nutrients such as EPA, DHA
and vitamin B12, have been commonly preferred as first choice diet for the rotifers [5].
* Corresponding author : stenlywullur@unsrat.ac.id© The Authors, published by EDP Sciences. This is an open access article distributed under the terms of the Creative Commons
Attribution License 4.0 (http://creativecommons.org/licenses/by/4.0/).

E3S Web of Conferences 147, 01005 (2020)
3rd ISMFR

https://doi.org/10.1051/e3sconf/202014701005

However, year round cultivation of sufficient and nutritious microalgae is a costly andlaborious task for many hatcheries all over the world, thus limits the continuous productionof sufficient live foods. Lack of sufficient live food sometimes disrupts the fish seedling
production programs in the microalgae-based hatcheries [6]. In our previous studies where
the FWD was used to culture different rotifer species, we obtained about 1,200 individuals
ml-1 of the rotifer, Brachionus rotundiformis [7], and about 1,600 individuals ml-1 of Proales
similis [8] within 10 days of culture. In addition, the rotifers had relatively higher EPA andDHA contents than those rotifers cultured using normal (non-enriched) C. vulgaris diet. Thisfinding demonstrates a possibility of obtaining nutrient-rich rotifers without necessarily
feeding them on enriched microalgae or supplementing them with the expensive commercial
emulsions. FWD therefore presents an opportunity to lower production cost in aquaculture
hatcheries. However, the suitability of the FWD-fed rotifers for larval fish rearing still
remains unclear. To explore on this aspect, the present study was conducted to determine theeffects of the rotifer, B. rotundiformis (SS-type) fed with FWD, on the larval growth anddevelopment of the Japanese whiting Sillago japonica. S. japonica is an ecologically valuable
fish resource distributed along the coasts of Japan, South Korea, Taiwan and the Philippines[9].

2 Materials and methods
2.1 Rotifer preparation
The inoculant rotifer, B. rotundiformis (SS-type; Thai strain) was obtained from Nagasaki
Prefectural Institute of Fisheries, Japan. The SS-type rotifer is small in size (i.e. 70 – 160µm), and is considered appropriate for the small-mouth fish larvae such as of S. japonica[10]. The rotifers employed in the test experiment were cultured using the FWD technologythat has been extensively described in our patent registered under number: P00201609066 inIndonesia. In this article, we have only presented a flow chart of the rotifer production unit
using FWD in Figure 1. Meanwhile, the rotifers used for the control experiment were batch
cultured in 50 l of artificial seawater (22 ppt) and fed with HUFA enriched Chlorella vulgaris
diet (Super Fresh Chlorella-V12, Chlorella Industry Co. Ltd., Fukuoka, Japan) at 25°C withaeration (Figure 2). The C. vulgaris diet was daily maintained at 7.0×106 cells ml-1 in therotifer cultures [11].
2.2 Rotifer swimming speed
To determine the viability of the rotifers, swimming speed was recorded for 30 seconds undera stereomicroscope at 10× (SteREO Discovery V8, Zeiss, Germany) equipped with a digital
camera (AxioCam, HSm) and image-analysis software (AxioVision 4.8). Five rotifers were
sampled from each treatment and placed in 20 µl water drop, in which the swimming speed
was measured using Dipp Motion Pro version 8.0 (DITECT Co. Ltd., Japan) in millimeters
per second (mm s -1).
2.3 Larviculture experimentFertilized eggs of S. japonica were obtained from the same institute as rotifers. The eggs were
carefully transferred into six polycarbonate tanks each containing 100 l of artificial sea water(33 ppt) at 10 eggs l-1 with aeration fixed at 50 ml min-1 [12]. After hatching, larval fish
development was monitored every 6 hours until mouth opening and then, the feeding process
began (Fig. 3). Two feeding regimes i.e. the test diet (FWD-fed rotifers) and control diet2

E3S Web of Conferences 147, 01005 (2020)
3rd ISMFR

https://doi.org/10.1051/e3sconf/202014701005(rotifers fed with enriched C. vulgaris) were each triplicated in the culture tanks. The fishlarvae were reared at 25°C with 12-h diurnal photoperiod (900–2100), for 10 days. The
density of rotifers (diet) in each larval fish rearing tank was maintained at 10 ind. ml -1
throughout the rearing period from 2 days post hatching (dph), which was the time of mouth
opening. Ten larval fish were randomly sampled every 2 days from 2 dph for morphometricand gut content analysis. The fish samples were kept in small screw-cap bottles anesthetizedwith 2-3 drops of MS 222 followed by 5% formalin fixation [12].

Fishwastes mainly heads (Fishwaste diet (FWD))

Crush FWD using mortar and pestle

Weigh 0.5 g/L of FWD and wrap in 200µm
plankton net and insert in about 150 cm2 text liner
container
Add 0.2 g/L of carbon
source (wheat flour)
water bath system
maintained at 28±1oC
Rotifer

1st harvesting day 5 andadd new water and FWD

FWD

2nd harvesting day 10 and
add new water and FWD

3rd harvesting day 14 andadd new water and FWD

Culture conditions
 Sea water 22ppt.
 inoculate 20
rotifers/mL
 Monitor water quality
in the tanks daily (NH3N, DO, and pH).

Wash and disinfect tanks
and restart cultures

Figure 1. Flowchart for rotifer production using the low-cost FWD technology

Figure 2. Rotifer batch cultures with HUFA enriched Chlorella vulgaris diet (Super Fresh

Chlorella-V12, Chlorella Industry Co. Ltd., Fukuoka, JapanThe hatching rate and survival activity index (SAI) experiments were performed todetermine the quality of the fish eggs and fish larval survivability during starvation,
respectively [13]. Here, 20 eggs were placed in a 500 ml beaker containing 300 ml of artificial
sea water (33 ppt) at 25°C in total darkness, without aeration and feeding. Dead larvae werecounted and removed every 24 h until total larval mortality was reached. Triplicate

3E3S Web of Conferences 147, 01005 (2020)
3rd ISMFR

https://doi.org/10.1051/e3sconf/202014701005

observations were used to calculate the hatching rate and SAI. The percentage of eggs that
hatched normally was calculated after 24 h by the formula: [Normally hatched larvae / total
number of eggs] × 100%, while SAI was calculated using the equation of [13],SAI = 1K N  hi  i
N

(1)

i 1

where N = total number of examined larvae, hi = cumulated mortality by i-th day and K =number of days elapsed until all larvae died due to starvation. Morphological characteristicsof the larval fish samples i.e. total and standard length, eye diameter, body depth and headlength (Fig. 3), were measured using a microscopic measurement system that included a
stereomicroscope (Discovery V8, Zeiss, Germany) equipped with a digital camera(AxioCam, HSm) and image-analysis software (AxioVision 4.8). At 10 dph, the percent
survival of the fish larvae was calculated from the average number of surviving fish larvaein each culture tank. Also, the viability test of the fish larvae was determined at 10 dph bysubjecting 10 fish larvae from each replicate tank to 30 s of air exposure before determining
the rate of their survival thereafter. The dry weight of the fish larvae was measured in preweighed aluminum boats dried at 60oC for 24 h and weighed using ultra-micro chemicalbalance (Mettler Toledo, USA).

Figure 3. Larval developmental stages of Sillago japonica; A) egg stage, B) newly hatched larvae
with a bigger egg yolk, C) 1 dph showing reduced egg yolk size, D) 2 dph showing mouth opening(end of exogenous feeding)

4

E3S Web of Conferences 147, 01005 (2020)
3rd ISMFR

https://doi.org/10.1051/e3sconf/202014701005

Figure 4. The five morphological characteristics to estimate larval growth and development of S.
japonica. TL, total length; SL, standard length; HL, head length; ED, eye diameter; BD, body depth

2.4 Fatty acid analysis
At 10 dph, all the fish larvae were harvested from each treatment tank, dried using filter paper
from beneath the harvesting mesh and kept at -80oC until biochemical analysis. The total
lipid and fatty acid composition of the fish larvae and rotifer samples were performed by
Chlorella Industry Co., Fukuoka, Japan. The sample methanol lysates were prepared at 100°C for 2 h after the addition of 2 M hydrogen chloride methanol. Fatty acid methyl esters(FAME) were extracted by n-hexane. Gas chromatography analysis was performed using a
GC-2010 (Shimadzu Scientific Instruments, Inc.) equipped with a HR-SS-10 column(Shinwa Chemical Industries, Ltd.). The column temperature was regulated at 150 to 220 °C.
Individual fatty acids were quantified by means of the response factor to 15:0 fatty acids as
the internal standard [14].
2.5 Statistical analysis
Statistical analysis was carried out using the R statistical software (version 3.2.1 of the R
Foundation for Statistical Computing Platform © 2015). The Bartlett test for homogeneity
was used to test the equality of variances. Repeated ANOVA measures were used to test theeffects of the diets on fish larval morphological parameters, gut content and swimming speed.
The larval fish survival and viability tests were analyzed using Kruskal-Wallis test and LogRank test for groups, respectively. Where significant differences were detected, the Tukey
HSD post hoc test was performed to locate them at p<0.05.

3 Results
The hatching rate of S. japonica eggs was 93.3±2.9%. The hatched fish larvae survived up to
5 days of starvation, with a SAI of 5.8±1.1. The rotifer swimming speed was affected by thediets (One way ANOVA, F=14.51, p=0.01), where the control-rotifers had significantly
higher speed (0.83 mm s-1) than FWD-rotifers (0.65 mm s-1) (Tukey HSD, p=0.01). The
number of ingested rotifers by individual fish larvae was significantly affected by age of fishlarvae (Two-way ANOVA, F=128.04, p=0.00), but not the diets (F=0.24, p=0.62). The fishlarvae from the two treatments ingested similar amounts of rotifers each day and, at 10 dph,
about 10.5±3.1 and 11.6±2.7 rotifers were ingested by the FWD-fish and control-fish,

5

E3S Web of Conferences 147, 01005 (2020)
3rd ISMFR

https://doi.org/10.1051/e3sconf/202014701005

respectively. At 10 dph, there was no significant differences in morphological parameters
e.g. head length (One-way ANOVA, F=2.46, p=0.13), eye diameter (One-way ANOVA,
F=2.46, p=0.12) and body depth (One-way ANOVA, F=0.16, p=0.69) between FWD-fishand control-fish. However, there was a significantly higher total and standard length for the
FWD-fish than the control-fish (p=0.00) (Table 1). There was no significant differences infish larval survival rate (Kruskal-Wallis test, χ2=0.19, df=1, p=0.66), viability (Log-rank testfor groups, χ2 = 0.4, df=1, p=0.55) and dry weight (One-way ANOVA, F=6.41, p=0.06)
between the two diets (Table 2). The nutritional value of rotifers and fish larvae issummarized in Table 3. Due to high fish mortality experienced during the experiment, thequantities of fish sample were insufficient for total lipid analysis. Nonetheless, percent
compositions of DHA were 5.2 and 18.2 for FWD- and control- fish, respectively. The EPA
was under detectable limit in both samples. The ratio of DHA: ARA was 2.4 and 8.3 for the
FWD- and control- fish, respectively (Table 4).
Table 1: Morphological parameters of the fish larvae as at 10 dph, the values are mean length (mm) ±SD, with different numbers of fish larvae per replicate. One-way ANOVA, Tukey HSD, different
superscripts in each column denote significant differences at p<0.05 between the diets, a>b

Diet
FWD
Control

Total length(n=30)
3.65 ± 0.30a
3.42 ± 0.22b

Standard length(n=30)
3.33 ± 0.33a
2.94 ± 0.33b

Head length(n=12)
0.76 ± 0.03
0.74 ± 0.05

Eye diameter(n=30)
0.25 ± 0.02
0.24 ± 0.02

Body depth(n=12)
0.69 ± 0.11
0.68 ± 0.11

Table 2: Means ± SD of fish larval characteristics at 10 dph. The fish dry weight at 2 dph was 0.08 ±0.01mg/ind in both diets, n=3

Diets
FWD
Control

Dry weight (mg/ind)
0.26 ± 0.03
0.21 ± 0.01

Survival rate (%)
9.70 ± 9.06
4.93 ± 0.83

Viability (%)
58.8 ± 9.3
64.4 ±7.9

4 Discussion
The results of this study show that fish larval rearing is possible with cheaply produced
rotifers. This is important for improving fish larviculture especially in the developing
countries, where aquaculture is a significant pillar for food and nutrition security. The fish
eggs employed in the experiment were of high quality (93.3±2.9% hatching rate) and, thefish larvae showed high survivability during starvation (SAI: 5.8±1.1). These results arecomparable to those of [15] who reported 95% and 4.1 – 8.4 hatching rate and SAI,
respectively for S. japonica larvae. There was unprecedented high fish mortality (i.e. survival
rates of 9.7±9.1 % and 4.9 ±0.8 % for FWD treatment and control, respectively), and thereason was not clearly determined. However, it was suspected that aeration rate employed in
the current study was probably not optimal for this fish species. It could have also beenpossible that the fish larvae suffered from unknown microbial influence. In our previous
study, FWD generated huge density of bacterial communities, of which some could possess
lethal characteristics to cultured animals [7,8]. Further studies are recommended to determine
the optimum aeration rate for S. japonica larvae, and also to screen the possible microbialcommunities contained in the culture facilities.
The viability of the FWD-fed rotifers as determined by the swimming speed may havebeen weaker than control-rotifers perhaps due to high bacterial turbidity in the FWD culture
medium. More active rotifers are normally preferred in larviculture because they improvelarval fish development due to the ability to stimulate fish raptorial behaviour [16].

6E3S Web of Conferences 147, 01005 (2020)
3rd ISMFR

https://doi.org/10.1051/e3sconf/202014701005

Table 3: Total lipids (Total, mg/g dry weight) and fatty acid composition (% of total lipids) of therotifers fed with FWD and control diet, and of the fish (S. japonica) larvae cultured with the FWDand control-rotifers for 10 days. * = total lipids not measured due to limited sample quantity

Rotifers fed with
FWD
Control dietS. japonica larvae fed with
FWD-fed rotifers
Control-rotifers

Total lipids

20.5

23.3

*

*

Fatty acids
14:0
14:1
16:0
16:1
16:2
18:0
18:1
18:2n-6
18:2n-3
20:0
20:1
20:4n-6
20:5n-3
22:0
22:1
24:0
24:1
22:5n-3
22:6n-3
Others
DHA/EPA
DHA/ARA
Total

5.4
2.3
15.2
3.0
0.6
4.3
4.8
4.1
0.5
0.3
0.6
0.0
1.9
0.0
15.1
1.4
0.8
0.5
1.7
37.4
0.9
0.0
100

3.6
0.2
18.7
2.1
1.2
7.0
14.7
1.3
0.7
0.8
0.3
2.5
6.6
0.3
0.4
1.3
0.5
2.2
18.8
16.8
2.8
7.5
100

1.1
0.4
14.5
1.1
1.3
10.0
4.9
21.8
2.4
0.0
0.3
2.2
0.0
1.0
2.7
0.4
1.1
3.0
5.2
26.6
0.0
2.4
100

2.8
0.1
17.4
5.1
0.5
5.6
13.6
9.5
1.0
0.4
0.1
2.2
0.0
0.3
5.0
0.6
0.3
2.1
18.2
15.3
0.0
8.3
100

The results showed that diet did not determine the number of rotifers ingested by thelarval fish, thus suggesting equal palatability between the FWD-fed rotifers and those fed onconventional enriched algal diet. Due to same ingestion rate of rotifers, the fish morphometricmeasurements (head length, eye diameter and body depth), survival, viability and dry weight.
However, a significantly higher total and standard length for the fish given FWD-rotifers thanthose given control diet was realized. These results indicate the superiority of FWD as rotifer
diet for fish larviculture production. The fact that there was no significant difference in survivalrate, viability, dry weight, gut content, head length, eye diameter, and body depth between the two diets
indicate that waste-fed rotifers are as good as established larviculture technologies.

Previous larval fish rearing data confirm that survivability of fish larvae strongly depends
on the amount of DHA and ARA present in the diet [17,18,19]. Since most marine fish larvae
cannot synthesize DHA from precursor molecules e.g. EPA or α-linolenic acid (ALA),
supplying DHA-rich feeds to fish larvae is important [16]. The DHA/ARA ratio of 2.4

7

E3S Web of Conferences 147, 01005 (2020)
3rd ISMFR

https://doi.org/10.1051/e3sconf/202014701005

obtained in this study is less than the optimal ratio of 10.0 required for proper larval fish
development [18]. Nonetheless, the similarity of larval fish development parameters between
the test and control diet suggests a possibility of FWD for the larviculture of S. japonica. The
technology for FWD-fed rotifers promises appears a major leap toward developing a cheapand nutritionally rich diet for the rotifers, which are valuable food sources in larviculture.
Based on the food value of the bioflocs [7, 8]. The FWD culture fluid can be considered acheaper enrichment medium for low quality larval fish foods e.g. bakers' yeast and, may
reduce the need for the expensive enrichment emulsions. However, further studies are needed
for this suggestion. Further experiments are also recommended to test the FWD in other fish
species for larviculture. The advantage of FWD is that it is significantly cheaper because
fishwaste, which is the main ingredient in the diet, can be obtained for free. Therefore, theFWD-rotifers provides the means to develop more economic ways of improving aquacultureof commercially important fish species in small-scale tropical hatcheries.
This research was partly supported by Japanese Society for the Promotion of Sciences Asia-Africa(24380108) and JSPS Kakenhi (17H03862) to Atsushi Hagiwara, as well as Japanese (Ministry of
Education, Culture, Sports, Science and Technology) MEXT program, which provided Ph.D.
fellowship to Erick O. Ogello in Nagasaki University, Japan.

References
1.

P. Lavens, P. Sorgeloos, Manual on the production and use of live food for aquaculture.
In: FAO Fisheries Technical Paper. No. 361. FAO, Rome (1996).
2. M. Yufera, M.J. Darias, The onset of exogenous feeding in marine fish larvae.
Aquaculture, 268, 53–63 (2007).
3. T. Watanabe, C. Kitajama, S. Fujita Nutritional values of live organisms used in Japanfor mass propagation of fish: a review. Aquaculture, 34, 115–143 (1983).
4. K. Hamre, A. Srivastava, I. RØnnestad, A. Mangor-Jensen, J. Stoss, Several
micronutrients in the rotifer Brachionus sp. may not fulfil the nutritional requirementsof marine fish larvae. Aquac Nutr, 14, 51–60 (2008).
5. I. Maruyama, T. Nakao, I. Shigeno, Y. Ando, K. Hitayama, Application of unicellularalgae Chlorella vulgaris for the mass culture of marine rotifer Brachionus. Hydrobiol,
358, 133–138 (1997).
6. E. Lubzens, O. Gibson, O. Zmora, A. Sukenik, Potential advantages of frozen algae(Nannochloropsis sp.) for rotifer (Brachionus plicatilis) culture. Aquaculture, 133,
295–309 (1995).
7. E.O. Ogello, S. Wullur, Y. Sakakura, A. Hagiwara, Composting fishwastes as low-costand stable diet for culturing Brachionus rotundiformis Tschugunoff (Rotifera):
influence on water quality and microbiota. Aquaculture, 486, 232–239 (2018).
8. N. Kagali, E.O. Ogello, Y. Sakakura, A. Hagiwara, Fish processing wastes as analternative diet for culturing the minute rotifer, Proales similis de Beauchamp. Aquac
Res, DOI: 10.1111/are.13707 (2018)
9. M. Kashiwagi, S. Kondo, W. Yoshida, M. Yoshioka, Effects of temperature and salinity
on hatching success of Japanese whiting Sillago japonica eggs. Suisanzoshoku, 48(4),637–642 (2000).
10. A. Araujo, A. Hagiwara, Screening methods for improving rotifer culture quality.
Hydrobiol, 546, 553‒558 (2005).

8

E3S Web of Conferences 147, 01005 (2020)
3rd ISMFR

https://doi.org/10.1051/e3sconf/202014701005

11. H-J. Kim, Y. Sakakura, I. Maruyama, T. Nakamura, K. Takiyama, H. Fujiki, A.
Hagiwara, Feeding effect of selenium enriched rotifers on larval growth anddevelopment in red sea bream Pagrus major. Aquaculture, 432, 273–277 (2014).
12. H. Shimma, A. Tsujigado, Some biochemical quality of bred scorpaenoid fish,
Sebastiscus marmoratus, and activities of their larvae. Bulletin of National ResearchInstitute of Aquaculture, 2, 11–20 (1981).
13. J. Folch, M. Lees, H.S. Stanley, A simple method for the isolation and purification of
total lipids from animal tissue. J Biol Chem. 226, 497–506 (1957).
14. Y. Oozeki, R. Hirano, Variations in egg qualities of the Japanese whiting, Sillagojaponica, in the spawning season, In Maclean JL, Dizon LB, Hosillos LV. (eds.,) The
First Asian Fisheries Forum. Asian Fisheries Society, Manila, Philippines p. 679–682(1986).
15. R. Masuda, T. Takeuchi, K. Tsukamoto, Y. Ishizaki, M. Kanematsu, K. Imaizumi,
Critical involvement of dietary docosahexaenoic acid in the ontogeny of schoolingbehaviour in the yellowtail. J Fish Biol., 53, 471–484 (1998).
16. J.R. Sargent, L.A. McEvoy, J.G. Bell, Requirements, presentation and sources ofpolyunsaturated fatty acids in marine fish larval feeds. Aquaculture, 155, 117–127(1997).
17. J.R. Sargent, L.A. McEvoy, A. Estevez, J.G. Bell, M.V. Bell, R.J. Henderson, D.R.
Tocher, Lipid nutrition of marine fish during early development: current status andfuture directions. Aquaculture, 179, 217–229 (1999).
18. H. Furuita, H. Tanaka, T. Yamamoto, M. Shiraishi, T. Takeuchi, Effects of n-3 HUFA
levels in broodstock diet on the reproductive performance and egg and larval quality of
the Japanese flounder, Paralichthys olivaceus. Aquaculture, 187, 387‒398 (2000).

9