Content uploaded by Graeme B Martin
Author content
All content in this area was uploaded by Graeme B Martin on Dec 12, 2015
Content may be subject to copyright.
Emu Farming
Reproductive Technology
A report for the Rural Industries Research
and Development Corporation
by Irek Malecki and Graeme Martin
March 2000
RIRDC Publication No 00/37
RIRDC Project No UWA-39A
ii
© 2000 Rural Industries Research and Development Corporation.
All rights reserved.
ISBN 0 642 58070 7
ISSN 1440-6845
Emu Farming – Reproductive Technology
Publication No. 00/37
Project No. UWA-39A
The views expressed and the conclusions reached in this publication are those of the author
and not necessarily those of persons consulted. RIRDC shall not be responsible in any way
whatsoever to any person who relies in whole or in part on the contents of this report.
This publication is copyright. However, RIRDC encourages wide dissemination of its research,
providing the Corporation is clearly acknowledged. For any other enquiries concerning
reproduction, contact the Publications Manager on phone 02 6272 3186.
Researchers’ Contact Details
Dr Irek Malecki
Faculty of Agriculture (Animal science)
University of Western Australia
Nedlands 6907
Western Australia
Phone: +61 8 93802518
Fax: +61 8 93801040
Email: imalecki@agric.uwa.edu.au
A/Prof. Graeme Martin
Faculty of Agriculture (Animal science)
University of Western Australia
Nedlands 6907
Western Australia
Phone: +61 8 93802528
Fax: +61 8 93801040
Email: gmartin@agric.uwa.edu.au
RIRDC Contact Details
Rural Industries Research and Development Corporation
Level 1, AMA House
42 Macquarie Street
BARTON ACT 2600
PO Box 4776
KINGSTON ACT 2604
Phone: 02 6272 4539
Fax: 02 6272 5877
Email: rirdc@rirdc.gov.au.
Website: http://www.rirdc.gov.au
Published in March 2000
Printed on environmentally friendly paper by Canprint
iii
Foreword
The emu is being farmed for the production of leather, meat and oil, but the future development of the
industry is limited unless the efficiency of production is improved and genetic progress accelerated.
This project addresses these issues by establishing a basis for development of reproductive technology
for the emu industry.
Under the current emu farming system, emus are mostly bred in monogamous pairs. Monogamous
mating has a direct cost in that it requires the retention of excessive males in the flock – birds that are
not otherwise productive yet consume feed and capital resources that could be used to manage more
females and therefore produce more eggs and growing chicks. However, more importantly,
monogamous mating almost completely arrests genetic improvement. Even without the constraints of
monogamy, natural mating can never achieve the male:female ratio that is possible with AI, so the
selection differential is minimal and most of the semen from valuable males is wasted. The birds are
not easy to transport, so multiple natural matings are only feasible within a single farm, and exchange
of desired genes is limited. Finally, the genes of superior birds or lines of birds cannot be preserved.
The obvious alternative to natural mating is the artificial insemination technology that has allowed
other industries to develop highly efficient lines for specific products or markets.
This publication reviews a series of experiments carried out over two breeding seasons on a selected
group of emus maintained at the UWA Emu Research Facility in Perth, Western Australia. It analyses
data collected in studies of the seasonality of semen production, quality and quantity, the short- and
long-term aspects of semen preservation, and the feasibility and efficiency of artificial insemination.
The results suggest that the artificial insemination is highly feasible for emu farming and that this
sunrise industry could easily adopt techniques for collection of semen, storage and preservation, and
artificial insemination.
This project was funded from RIRDC Core Funds which are provided by the Federal Government.
This report, a new addition to RIRDC’s diverse range of over 450 research publications, forms part of
our New Animal Products R&D program, which aims to accelerate the development of viable new
animal industries.
Most of our publications are available for viewing, downloading or purchasing online through our
website:
• downloads at www.rirdc.gov.au/reports/Index.htm
• purchases at www.rirdc.gov.au/pub/cat/contents.html
Peter Core
Managing Director
Rural Industries Research and Development Corporation
iv
Acknowledgements
We would like to thank Dr Anne Jequier for donation of the programmable freezer and the liquid
nitrogen tank that was essential for the liquid preservation of emu semen.
Dr Gerard Smith and Ms Maeve Harvey from the Animal Health Laboratories, Agriculture Western
Australia, for their help in determined the fatty acid profiles of emu spermatozoa.
Dr James O’Shea from the Department of Zoology, University of Western Australia, kindly lent us the
fluorescence microscope.
v
Contents
Foreword.................................................................................................................................iii
Acknowledgements.................................................................................................................iv
List of Tables...........................................................................................................................vi
Executive Summary ...............................................................................................................vii
1. Introduction...........................................................................................................................1
2. Objectives.............................................................................................................................3
3. Methodology.........................................................................................................................3
4. Results .................................................................................................................................6
5. Discussion..........................................................................................................................28
6. Implications ........................................................................................................................30
7. Recommendations .............................................................................................................31
8. Bibliography........................................................................................................................32
vi
List of Tables
Table 1 Proportions of live and damaged sperm as detected by dual fluorescence and
nigrosine-eosin stains in fresh emu semen (values are means of 2 replicates).........6
Table 2 Effect of season on mean ejaculate characteristics...................................................9
Table 3 Effect of season on the concentration of major inorganic ions in emu seminal
plasma......................................................................................................................11
Table 4 Fatty acid content of the emu spermatozoa. ............................................................11
Table 5 Effect of season on the concentrations of major saturated and unsaturated fatty
acids.........................................................................................................................13
Table 6 Concentrations of major inorganic ions in the emu seminal plasma and comparison
with chicken and turkey seminal plasma (values are mM/L)....................................13
Table 7 The composition of semen diluents..........................................................................14
Table 8 Characteristics of diluents that were investigated, in relation to emu
seminal plasma. .......................................................................................................18
Table 9 Median duration of fertility following a single insemination with fresh semen. .........23
Table 10 Composition of cryopreservation diluent (Wishart, 1995).........................................23
Table 11 Osmotic conditions at different concentrations of cryoprotectant.............................25
List of Figures
Figure 1 Effect of season on the production of semen and spermatozoa in the emu. .............8
Figure 2 Effect of diluent and storage time on the number of live spermatozoa. ...................14
Figure 3 Effect of diluent and temperature on the number of live emu spermatozoa.............15
Figure 4 Effect of diluent and storage time on the number of live emu spermatozoa
stored at 4°C. ...........................................................................................................15
Figure 5 Effect of diluent and storage time on the number of live spermatozoa
stored at 20°C. .........................................................................................................16
Figure 6 Effect of diluent and storage time on the number of live spermatozoa
stored at 39°C. .........................................................................................................16
Figure 7 Effect of diluent and temperature on motility of emu spermatozoa. .........................17
Figure 8 Effect of diluent and storage time on motility of emu spermatozoa..........................17
Figure 9 Effect of dilution and diluent on motility of emu spermatozoa. .................................18
Figure 10 Effect of diluents on the number of live spermatozoa stored at 20°C......................19
Figure 11 Effect of diluents on the motility of emu spermatozoa stored at 20°C......................19
Figure 12 Effect of cryoprotectant on the integrity of spermatozoal membrane.......................24
Figure 13 Effect of cryoprotectant concentration on motility and number of live
emu spermatozoa following freezing........................................................................26
Figure 14 Effect of freezing protocol and cryoprotectant on the number of live emu
spermatozoa.............................................................................................................27
vii
Executive Summary
The emu is being farmed for the production of leather, meat and oil, but the future development of the
industry is limited unless the efficiency of production is improved and genetic progress accelerated.
Under current farming system, genetic improvement is almost completely arrested because emus are
bred in pairs. Monogamous mating has lead to retention of excessive males in the flock that are not
otherwise productive yet consume feed and capital resources that could be used to manage more
females and therefore produce more eggs and growing chicks. A potential solution to all of these
issues is artificial insemination (AI).
The development of AI technology for any industry involves development of techniques for semen
collection, systems for rearing and managing males for high semen yields, technologies for
preservation of semen and artificial insemination, and systems for the management of females for high
egg production and fertility. Other bird industries, including those based on the turkey, chicken, duck,
goose and guinea-fowl, make extensive use of artificial insemination. Perhaps the best comparison
with the emu is the guinea fowl, for which AI was introduced to reduce the cost of housing and
feeding the males because these birds are also virtually monogamous.
High yields of semen and spermatozoa throughout the entire period of egg production are critical in AI
systems. The major constraint to high and sustained sperm production by the emu is seasonality. Our
studies on the annual cycles in reproductive hormones and testicular function in the emu suggested
that the onset of the breeding season (beginning of sperm production) approaches slowly but its
cessation (termination of sperm production) is rather abrupt. These two critical periods will have an
impact on the quantity, fertilising ability of spermatozoa and therefore availability of spermatozoa for
AI. The number of females that can be inseminated from one ejaculate depends on the number of
spermatozoa required per dose and the frequency of insemination needed for maintenance of
maximum fertility. Female emus can lay fertilized eggs for up to three weeks following AI or natural
mating, but our early indications were that this is highly variable so that maximum fertility may
require weekly inseminations. Developing good insemination technique and determining the minimum
dose of fresh spermatozoa would let us know what are the requirements for AI in the emu.
Storage and dilution of semen require a good diluent, an essential component in any AI system. For
short term storage, the period of time between collection and deposition in the female reproductive
tract, the optimal temperature and diluent are needed as well as maximum semen/diluent dilution ratio
if we are to utilise the semen of a superior sire to the maximum and to be able to transport genes
between farms. There is no universal diluent for avian semen due to species differences in the
composition of seminal plasma, sperm enzyme systems, and metabolic requirements of sperm. It is
likely, therefore, that emu semen would store better at low temperatures but the best range needs to be
determined. We have no knowledge of the ionic content of emu seminal plasma, another factor that is
critical in diluent design. The only choice is to develop an emu semen diluent and test a range of
storage conditions. In addition, an indefinite storage would allow the banking of superior genes and
their transport over the long distances that characterize the Australian industry. It could also provide a
reservoir of spermatozoa during periods of lower sperm production. This requires development of the
protocol for freezing emu semen, but the best cryoprotectant and its concentration that has not yet
been identified.
The project aimed to establish a basis for development of reproductive technology for the emu
industry. A series of experiments was carried over two breeding seasons on a selected group of emus
maintained at the UWA research facility in Perth, Western Australia.
The production of semen and spermatozoa is very seasonal, lasts for about 6 months and can be
divided into 3 periods: the beginning, middle and the end. It appears that it takes more time for males
to attain the peak of production at beginning than it takes for them to decline from the peak to its
termination. Semen can be collected as early as March, which is about when female emus start egg
laying, but the libido of males is not as high as in the middle of the breeding season and initially
semen can only be collected once a day. The output of spermatozoa is also lower than for the rest of
viii
the season as the concentration of semen is low. As the production of semen and spermatozoa reaches
its peak in June-July, more spermatozoa are available for AI, however, the production of spermatozoa
and their quality starts decreasing soon after. This time of the breeding may be additionally affected by
females having lower ability to retain spermatozoa and also by some males stopping their semen
production.
Good storage and preservation techniques are essential in any AI program. Our emu semen diluent
needs development but the results are very encouraging. Because the levels of potassium, sodium,
calcium and magnesium in emu seminal plasma are similar to those in the chicken and turkey, we
anticipated that commercial poultry diluents should be suitable for storing emu semen. Our studies
clearly show that only Lake’s diluent can be recommended. Emu semen can be stored in Lake’s
diluent at either 4 or 20°C for up to 6 hours without the loss of viability. Storage in the newly
developed emu diluents should result in more live spermatozoa. Our diluents (UWA-E1, UWA-E2
and UWA-E3) maintained more viable spermatozoa than any other diluent and our results indicate that
sperm viability can be extended by reducing motility and that this could be achieved by increasing the
concentration of potassium and reducing sodium in the diluent.
Emu spermatozoa can be frozen in liquid nitrogen. They have low susceptibility to cold shock and a
slow cooling/freezing rate is most efficient. The recovery of viable spermatozoa (40%) is as good as in
the chicken. The survival of spermatozoa might be improved further by genetic selection and by
controlling the nutrition of the donor males.
Deposition of spermatozoa in the female oviduct (AI) can be successful and efficient if there is good
cooperation between the female and the inseminator. The crouching behaviour that female emus
develop allows for a stress-free approach, that would not interfere with ovulation. The insemination
technique can be learned and used, and little intervention in the cloaca is needed because the non-
speculum technique has also proven to have potential.
An elite male producing average ejaculates could be used to fertilise 32 females every 10 days, or up
to 320 females in a season. We know, however, that the output of spermatozoa is seasonal and the
male:female ratio will vary depending on the period of breeding season. However, as there is
considerable variation in sperm production between males, high sperm producers could be selected
and the male:female ratio could be increased further. Artificial insemination could also be made more
efficient by storing semen in the right diluent and by selecting females that show a longer duration of
fertility.
In summary, we have demonstrated that emus can be bred very effectively by artificial insemination.
Good quality ejaculates can be collected from males into the artificial cloaca by two methods. Using
these methods, male emus can be trained and their sperm production potential assessed. Similarly,
female emus can be selected for the breeding program based on their egg laying records and their
crouching behaviour. The emu diluent is yet to be fully developed, but spermatozoa for artificial
insemination can be stored successfully for up to 6 hours at 4 or 20°C in Lake’s diluent diluted 1:2.
Finally, emu spermatozoa can be cryopreserved for sperm banking or for transport over long distances.
At present, the best recovery is achieved by freezing pre-cooled (to 5°C) sperm samples containing 9%
DMA at the rate of 1°C/min to –35°C, and then plunging them into liquid nitrogen. Once we have
introduced proper selection programs on farms, the efficiency of production should increase and this
would further benefit the producers.
In conclusion, by introducing reproductive technology, the emu industry can now make use of the
massive reservoir of elite genes and genetic diversity at our disposal. The artificial insemination can
substitute for natural mating and greatly reduce the male to female ratio. As season affects the
availability of spermatozoa for AI, early and late sperm producing males could be sought to meet these
demands. Males that produce sperm from March until September would be best, and we might be able
to select them from existing flocks, or breed selectively for longer duration of sperm production. This
is one of many breeding objectives that emu farming can now focus on with this technology.
ix
It needs to be stressed that the results of this study come from a select group of trained birds
maintained at the UWA research facility. These birds, as with those on commercial farms, have not
been subjected to much selection pressure and, as the variation between individuals suggests, there is a
lot of room for genetic selection in a number of traits, such as semen output, libido, egg production
and fertility.
The fertilising ability of emu spermatozoa following storage and cryopreservation needs to be
examined further because it is overestimated by the conventional techniques (sperm cell integrity and
motility) that we used. Fertility trials would be the most accurate method to study this, but it is very
slow and costly. A new approach using a sperm-egg interaction assay could be developed, and this
would not require the expensive and time-consuming AI trials with large numbers of females, daily
egg collection and time-consuming incubation. The sperm-egg interaction assay could also be applied
to assess fertility of females. The length of their fertile period and the efficiency of sperm transport in
the oviduct could be determined by counting sperm trapped in the outer perivitelline layer of laid eggs.
Furthermore, the sperm egg interaction assay could be used to assess fertility of flocks.
Finally, now that we have established this technology for the emu industry, and we have new
directions for greatly improving it, it is also time to consider transferring the concepts to the ostrich
industry.
1
1. Introduction
The development of AI technology for any bird industry involves development of techniques for
semen collection, systems for rearing and managing males for high semen yields, technologies for
preservation of semen and artificial insemination, and systems for the management of females for high
egg production and fertility. Other bird industries, including those based on the turkey, chicken, duck,
goose and guinea-fowl, make extensive use of artificial insemination (Lake & Stewart, 1978). A very
good comparison with the emu is the guinea fowl, for which AI was introduced to reduce the cost of
housing and feeding the males because these birds are also virtually monogamous (Etches, 1996).
High yields of semen and spermatozoa throughout the entire period of egg production are critical in AI
systems. The major constraint to high and sustained sperm production by the emu is seasonality . Our
studies on the annual cycles in reproductive hormones and testicular function in the emu suggested
that the onset of the breeding season approaches slowly but terminates rather abruptly. The pituitary-
gonadal axis appears to be activated as early as January, but the testes do not attain full activity until
May-June. Semen collection studies that began in mid-season have shown that the output of
spermatozoa gradually declines (Malecki, et al. 1997b) and then terminates in September-October.
Because the testes undergo a complete cycle of growth and regression every year, we anticipated that
sperm production have a phase of acceleration at the beginning of the season, and cessation at the end
of it. The impact of these two critical periods on the quantity and fertilising ability of spermatozoa was
not known.
The number of females that can be inseminated from one ejaculate will depend on the number of
spermatozoa required per dose and the frequency of insemination needed for maintenance of
maximum fertility. Poultry species are generally inseminated at weekly intervals regardless of the
length of their fertile period, since extending the insemination interval reduces fertility (Brillard,
1994). Female emus can lay fertile eggs for up to three weeks following AI or natural mating, but
early indications were that this is highly variable so that maximum fertility may require weekly
inseminations (Malecki et al. 1996). Developing good insemination techniques and determining the
minimum dose of fresh spermatozoa would allow us to decide on the requirements for AI in the emu.
Storage and dilution of semen require a good diluent, an essential component of any AI system. For
short term storage, the period of time between collection and deposition in the female reproductive
tract, we need to know the optimal temperature and the best diluent composition, as well as the
optimal semen:diluent dilution ratio for extension, if we are to utilise the semen of a superior sire to
the maximum, and be able to transport his genes. There is no universal diluent for avian semen due to
species differences in the composition of seminal plasma, sperm enzyme systems, and metabolic
requirements of sperm (Lake & Wishart, 1984; Wishart, 1989). Fowl semen stores better at low
temperatures (5-15°C) so it is likely that emu semen would be the same, but the optimal range needs to
be determined. Spermatozoa also require energy in the form of carbohydrates (glucose or fructose) to
maintain their motility, and semen clearly benefits from adding this exogenous source of energy into
the storage medium (Lake, 1960; Sexton & Fewlass, 1978).
The ionic composition of the diluent is usually based on the composition of seminal plasma (Lake,
1960; Lake & Wishart, 1984). However, depending on the storage temperature, the optimal
concentration of major ions or cations may be different (Chaudhuri & Lake, 1988) and completely
different diluents may be equally useful. We had no knowledge of the ionic content of emu seminal
plasma so the only choice was to study seminal plasma, develop an emu semen diluent and test a range
of storage conditions. In addition, indefinite storage would be useful because it would allow the
banking of superior genes and their transport over the long distances that characterize the Australian
industry. It could also provide a reservoir of spermatozoa during periods of lower sperm production.
Protocols for freezing avian semen have been developed and, while they are successful for the
chicken, results are variable with other species (Lake & Stewart, 1978b; Lake, 1986). Glycerol,
dimethylsulphoxide and dimethylacetamine have been commonly used as cryoprotectants, although
the list of possibilities is much longer. A major problem is toxicity of cryoprotectant, but this depends
on the concentration of the cryoprotectant and the composition of the semen diluent (Lake & Ravie,
2
1984; Sexton, 1980). To our knowledge, none had been tested with emu semen, so we needed to
determine the best cryoprotectant and the optimal concentration.
3
2. Objectives
• To determine the capacity of emus to produce sperm;
• To develop protocols for preservation of emu semen;
• To develop methods of artificial insemination for the emu industry.
3. Methodology
1. Semen collection
Semen was collected into an artificial cloaca from 6 males. Three males have been trained for
collection using teaser, and the other three trained without teasers (Malecki et al. 1997b).
2. Evaluation of semen
2.1.Concentration of spermatozoa
Following collection, 20 µl of semen was diluted 1:400 in 0.9% NaCl solution and formaldehyde and
the absorbance read on a Shimadzu spectrophotometer. Concentration of spermatozoa was then
determined from the standard curve. Prior to that, the optimum wavelength (550 nm) and the standard
curve had been estimated. The concentration of spermatozoa was determined by counting sperm on the
haemocytometer and plotted against the absorbance of the same sperm suspension.
2.2.pH
Semen pH was determined immediately after collection with the MC-80 pH-mV-Temperature Meter
(TPS, Brisbane, Australia) by placing the electrode into the collecting tube containing semen.
2.3.Osmolarity
Semen and seminal plasma osmolarity were determined with the Fiske ONE TEN Osmometer (Fiske
Associates, Needham Hights, MA 02194, USA).
2.4. Proportions of live and normal spermatozoa
The nigrosine-eosin staining technique was used (Lake & Stewart, 1978). A 10 µl aliquot of semen
was placed on a microscope slide, then 50µl nigrosine-eosin stain mixture was added, gently stirred
and smeared over the slide. Any excess of semen and stained mixture was removed and the smear was
dried with a hair drier. Proportions of live and normal spermatozoa were determined by counting a
minimum of 200 spermatozoa on the slide.
2.5. Motility
Semen was given an arbitrary score from 0 - 5 (Allen & Champion 1955) based on the following
assessment:
0 0%, no motility discernable;
1 1 – 20% of sperm exhibiting slight undulating movement; mostly weak and oscillatory;
2 20 – 40% of sperm showing undulatory movement; no waves or eddies formed; there maybe a
number of inactive sperm;
3 40 – 60% of sperm showing progressive motility; vigorous motion; slowly moving waves and
eddies produced;
4 60 – 80% of sperm showing progressive motility; waves and eddies of great rapidity of formation
and movement;
4
5 80 – 100% of sperm in vigorous and progressive movement; extremely rapid formation of eddies
and movement.
5
3. Determination of ions in seminal plasma
The seminal plasma was separated from spermatozoa by double centrifugation at 200 xg for 10 min at
20°C and then stored at –20°C. The concentrations of potassium, sodium, calcium and magnesium
were determined by atomic absorption spectrophotometry.
4. Fatty acid profiles
Spermatozoa separated from seminal plasma following double centrifugation was frozen and stored at
–20°C until assayed by the high performance liquid chromatography (HPLC).
5. Cryopreservation of spermatozoa
Emu semen was collected, pooled and then diluted 1:1 with cryopreservation diluent. The temperature
of the samples was reduced to 5°C by holding the samples in the –20°C freezer. Three cryoprotectants
were investigated: glycerol (GLY), dimethylsuphoxide (DMSO) and dimethylacetamide (DMA) at
concentration of 3, 6, 9 and 12% of cryoprotectant in diluted semen. The programmable freezer was
used to freeze semen according to entered protocols.
6. Artificial insemination
The females were inseminated intravaginally. The inseminator followed the female until she assumed
the crouched mating position. The insemination straw was then inserted into the vagina with the aid of
a speculum and the semen was injected as the straw was being slowly withdrawn.
7. Egg collection, storage and incubation
The eggs were collected early in the afternoon soon after they were layed, but if they were layed late
in the evening they were collected the next day in the morning. They were then stored for up to 3 days
at 15°C and then incubated at 36.5°C for up to 3 weeks in the emu egg incubator.
8. Egg fertility
The eggs were candled on Days 7 and 14 of incubation using the emu egg candler. The eggs that were
identified as clear were broken open and examined for sign of embryonic development. The eggs
without sign of development was declared infertile.
Statistical analyses of data
Data were analysed by ANOVA. Effect of season on semen quality and quantity was analysed by the
repeated measures model. Factorial analyses were used to determine the effect of storage and freezing
on the number of live and motility of spermatozoa.
6
4. Results
Experiment 1: Evaluation of the methods for estimating viability of emu spermatozoa
Evaluation of the methods for estimating viability of emu spermatozoa was needed to determine which
staining technique would best detect and differentiate between live, dead and damaged cells, in either
fresh or stored emu spermatozoa. Initial attempts to evaluate several techniques had to be modified
due to time constraints, availability of equipment and technical application of the results. A flow
cytometer could not be borrowed for the laboratory in Shenton Park, so this technique had to be
excluded. The nigrosine-eosin-Giemsa technique was not applicable to emu spermatozoa. because this
combination of stains could not differentiate between sperm with damaged and undamaged plasma
membranes, perhaps due to the morphological differences between avian and mammalian
spermatozoa. We therefore compared nigrosine-eosin and fluorescence stains (SYBR-14, Propidium
Iodide and Calcein-AM) for the detection of viable sperm in fresh emu semen. Propidium iodide can
only enter damaged cells, in which it binds to and stains cellular DNA red. SYBR-14 and Calcein-AM
stain viable sperm green. Nigrosine-eosin and fluorescence stains detected similar proportions of
viable sperm (Table 1).
Table 1. Proportions of live and damaged sperm as detected by dual fluorescence and nigrosine-eosin
stains in fresh emu semen (values are means of 2 replicates).
Actual % Live and Damaged Spermatozoa
N-E SYBR + PI CAL + PI CAL + PI SYBR + PI
Theoretical %
Live Sperm White* Green Green Dual Dual
0 0.0 0.0 0.0 0.0 0.0
20 20.0 28.5 26.5 0.6 0.0
40 47.7 49.0 43.4 1.6 1.1
60 68.0 59.3 57.8 2.1 1.5
80 71.4 75.0 61.6 2.9 0.9
100 86.6 92.2 76.7 2.4 0.7
*Legend
N-E: combination of nigrosine and eosin stains
SYBR + PI: combination of SYBR-14 (SYBR) and PI (propidium iodide) stains
CAL + PI: combination of CAL (Calcein-AM) and PI (propidium iodide) stains
White: live and undamaged sperm remain unstained as opposed to dead sperm assuming pink colour
Green: colour assumed by live sperm while dead sperm become red
Dual: both colours red and green
Viable sperm that were stained with any of the 3 combinations of stains correlated (P < 0.001) strongly
with the theoretical percentage of viable sperm (N-E, r = 0.97; SYBR, r = 0.98; Calcein, r = 0.96),
indicating that all 3 stain combinations could be used to assess fresh sperm. However, when
morphological examination of sperm is needed, the nigrosine-eosin stain is the only choice as the
fluorescence stains allow the examination of live cells and are not stable and fade relatively fast when
exposed to ultraviolet light. When using nigrosine-eosin stains, semen smears can be preserved and the
morphology of the sperm can be studied at any time. We therefore planned to assess the use of
fluorescence stains in the stored and cryopreserved sperm as they have been reported to be better than
the nigrosine-eosin stains (Chalah & Brillard, 1998). Unfortunately, the fluorescence microscope
could not be borrowed again in the 1999 breeding season so this investigation has been delayed. We
therefore report on the use of nigrosine-eosin stains in the experiments that required the examination
of sperm viability or morphology.
7
Experiment 2: Effect of season on the quantity and quality of emu semen and spermatozoa
This investigation began in July 1998 and was completed in July 1999. Six males were trained to
ejaculate into the artificial cloaca and their semen was collected twice a day (09.00 h and 15.00 h),
twice a week. Semen volume, sperm concentration, number of spermatozoa per ejaculate and pH of
semen were measured. Nigrosine-eosin smears were prepared for each ejaculate to determine the
percentage of live and normal spermatozoa under light microscopy. Seminal plasma was separated
from spermatozoa by centrifugation. Spermatozoa were used to determine the fatty acid profile while
the seminal plasma was assayed for inorganic ions.
Effect of season on the quantity of emu semen and spermatozoa (semen volume, concentration and
total number of spermatozoa)
The production of semen and spermatozoa was very seasonal, beginning in March and terminating in
September (Fig. 1). The males were already sexually active in mid-February and 2 males could be
induced to mate but their ejaculates contained few spermatozoa. Collection of semen began in March
but the libido of males was low and they could only ejaculate once a day. The production of semen
and spermatozoa increased steadily reaching mid-season values in May, but the peak of production
was in June and July. Then the production started decreasing gradually. Some males stopped semen
production in August and the others in September.
8
0
1
2
3
4
Number of spermatozoa (x109)
0
1
2
3
4
Sperm concentration (x109/ml)
0
0.5
1
Semen volume (ml)
March - April - May
June - July
August - September
Figure 1. Effect of season on the production of semen and spermatozoa in the emu.
The mean ejaculate (± se, n = 294) had a volume of 0.64 ± 0.0 mL, contained 2.0 ± 0.1 x109 sperm
and had a concentration of 2.9 ± 0.1 x 109 sperm/mL. The mean volume, concentration and number of
spermatozoa differed between times within the breeding season (Table 2). The mean ejaculate volume
(range, 0.35 – 1.25 mL), concentration (range, 1.52 – 4.15 x109 sperm/mL) and number of
spermatozoa (range, 0.69 – 4.48 x109) differed between males (P<0.01).
9
Table 2. Effect of season on mean ejaculate characteristics.
Part of Breeding Season
Parameter
March - April –
May
Beginning of Season
Jun – July
Mid-Season
August – September
End of Season
Volume (mL)
Concentration (x109/mL)
Number of spermatozoa
(x109)
Live spermatozoa (%)
Abnormal spermatozoa (%)
0.54 ± 0.0a
1.9 ± 0.1b
1.1 ± 0.1a
92.8 ± 0.8a
3.1 ± 0.2
0.90 ± 0.1b
3.4 ± 0.1a
3.1 ± 0.3b
92.1 ± 0.7a
3.0 ± 0.3
0.61 ± 0.0a
3.4 ± 0.1a
2.3 ± 0.2c
86.7 ± 0.1b
3.3 ± 0.3
*Values with different superscript within rows differ significantly (P<0.05)
Effect of season on the quality of emu semen and spermatozoa (concentrations of potassium, sodium,
calcium, magnesium, pH, fatty acid profiles of spermatozoa, proportions of live and abnormal
spermatozoa)
Because emus breed seasonally, we anticipated that concentrations of ions in the seminal plasma
would be affected by season. The ionic composition of the diluent is usually based on the composition
of seminal plasma so we needed to establish the extent to which season could affect seminal plasma
and whether such changes would require any adjustments in the diluent composition.
The pH of semen can affect the motility of spermatozoa. The pH of emu semen was not known but
was necessary in order to formulate a diluent of the correct pH. We also needed to know if the pH is
affected by season.
The fatty acid composition of spermatozoa depends on nutrition and age. In addition, the type and
concentrations of fatty acids can affect the susceptibility of spermatozoa to low temperatures. The
fatty acid composition of emu spermatozoa was thus investigated to determine the type of fatty acids
present, the ratio of unsaturated to saturated fatty acids, and the effect of season.
We have previously determined mean number of live and abnormal spermatozoa in the emu ejaculate,
but the effect of season is not known and needs be determined because it will affect the number of live
and normal spermatozoa available for artificial insemination.
Inorganic ions
The concentrations of potassium and sodium were affected by season (Table 3). High potassium levels
were in middle of the season and low levels at the beginning and end of the season. Sodium was high
from the beginning of breeding season to mid-season and low at the end of the season. Calcium and
magnesium levels did not change throughout the season. The mean concentrations of potassium,
calcium and magnesium in seminal plasma differed between males (P < 0.05).
pH
The mean pH of semen was 7.4 ± 0.1 and was not affected by season. The pH levels did not differ
between males and their mean values ranged from 7.1 to 7.5.
10
11
Table 3. Effect of season on the concentration of major inorganic ions in emu seminal plasma
(values are mM/L).
Part of Breeding Season
Inorganic ion March - April – May
Beginning of Season
Jun – July
Mid-Season
August – September
End of Season
K+
Na+
Ca+2
Mg+2
13.6 ± 0.3a
135.5 ± 0.8d
2.1 ± 0.0
1.5 ± 0.1
17.7 ± 0.4b
131.3 ± 0.1d
2.2 ± 0.0
1.5 ± 0.1
13.8 ± 0.4a
119.5 ± 0.2c
2.0 ± 0.1
1.7 ± 0.3
*Values with different superscript within rows differ significantly (P<0.05)
Live and abnormal spermatozoa
The emu ejaculate contained an average of 90.0 ± 0.5% (range 33.0 – 100.0) of live and 3.1 ± 0.1%
(range 0.0 – 30.0) of abnormal spermatozoa. The number of live spermatozoa was lower at the end of
the season compared to the middle and the beginning of the season, but the number of abnormal
spermatozoa was not affected by season (Table 2). Thus, less live spermatozoa will be available for AI
or storage at the end of the season and, but on average, the emu ejaculate contains only 90% live
sperm, a problem that needs to be considered in preparing insemination doses.5
Fatty acids in emu spermatozoa
Emu spermatozoa contain a range of short- and long-chain saturated and unsaturated fatty acids (Table
4). The major saturated fatty acids were palmitic and stearic acids which accounted for nearly 40% of
the total. The major unsaturated fatty acids were arachidonic, docosatetraenoic and heneicosanoic
acids. Three fatty acids could not be determined and they accounted for about 10% of the total fatty
acids.
In emu spermatozoa, the ratio of unsaturated to saturated fatty acids, based on the 90% that are
known, is 1.23. The spermatozoa of the bull, ram and boar have higher ratios (2.5 – 3.0) and are
known to be more susceptible to cold shock damage, whereas rabbit, human and fowl spermatozoa
have a ratio of about 1 and they are less susceptible to cold shock damage. The estimated ratio in emu
spermatozoa indicates that they might have low susceptibility to cold shock damage during chilled
storage or cryopreservation. This is very encouraging for the future of reproductive technology in this
industry.
Table 4. Fatty acid content of the emu spermatozoa.
Fatty acid Carbons % Fatty acid Carbons %
Unknown UNK1 5.34 ± .2 Heneicosanoic C 20:1 7.1 ± 0.3
Palmitic C 16:0 16.7 ± 0.3 Eicosadienoic C 20:2 2.6 ± 0.3
Palmitoleic C 16:1 0.21 ± 0.1 Unknown UNK3 1.6 ± 0.2
cis-10-
Heptadecanoic C 17:1 0.28 ± 0.1 Arachidonic C 20:4 14.7 ± 0.5
Unknown UNK2 4.5 ± 0.3 Docosenoic C 22:1 0.36 ± 0.2
Stearic C 18:0 22.9 ± 0.5 Docosatetraenoic C 22:4 13.3 ± 2.0
Oleic C 18:1 8.7 ± 0.3 Tetracosanoic C 24:0 0.45 ± 0.2
Linoleic C 18:2 1.8 ± 0.1 Docosahexaenoic C 22:6 0.27 ± 0.1
The concentrations of the major saturated and unsaturated fatty acids were affected by season and the
values were lower at the end than the beginning of the season, except for docosatetraenoic acid which
12
was unexpectedly high (Table 5). We will re-examine this acid again in the 2000 breeding season.
Seasonal changes such as these could lead to increased sensitivity of spermatozoa to low temperatures,
and thus a poor outcome for storage. This needs investigation.
13
Table 5. Effect of season on the concentrations of major saturated and unsaturated fatty acids.
Part of Breeding Season
Fatty acid March - April – May
Beginning of Season
Jun – July
Mid-Season
August – September
End of Season
Stearic (C18:0)
Oleic (C18:1)
Linoleic (C18:2)
Arachidonic (C20:4)
Docosatetraenoic
(C22:4)
24.2 ± 0.5a
9.6 ± 0.5b
2.1 ± 0.1b
15.0 ± 0.6a
9.7 ± 0.3a
22.8 ± 0.3b
8.1 ± 0.4a
1.7 ± 0.1a
15.4 ± 0.7a
11.0 ± 0.4b
20.4 ± 0.3c
8.3 ± 0.5a
1.7 ± 0.1a
12.9 ± 1.3b
25.1 ± 1.2 c
*Values with different superscript within rows differ significantly (P<0.01)
Experiment 3: Composition of emu seminal plasma
Using semen samples obtained in Experiment 2, we determined the major inorganic ions. The levels of
potassium sodium, magnesium and calcium were similar to those reported for the chicken and turkey
(Table 6). This results suggest that both chicken and turkey semen diluents should be suitable for
storing emu semen. These data were considered in formulating emu semen diluents.
Table 6. Concentrations of major inorganic ions in the emu seminal plasma and comparison with
chicken and turkey seminal plasma (values are mM/L).
Major
inorganic ions Emu Chicken Turkey
K+
Na+
Ca+2
Mg+2
15.0 ± 0.2
130.0 ± 0.8
2.1 ± 0.3
1.58 ± 0.1
13
145
2.3
1.4
17.0; 17.9
136.0; 167.0
4.3
0.3; 1.0
Experiment 4: Effect of diluent components, dilution rate and temperature on the viability of
emu spermatozoa stored in vitro
In a series of experiments, we tested the effect of storage temperature, energy content, and ionic
composition on sperm survival. We tested 3 temperatures (4, 20, 39°C) and 3 dilution rates (1:2, 1:4,
1:8). Following collection, semen was pooled and diluted with either Lake’s, BPSE, Phosphate and
NaCl-TES diluents (Table 7), or left undiluted as a control. We measured motility (arbitrary score 0-5)
and the proportion of live spermatozoa using nigrosine-eosin stains.
14
Table 7. The composition of semen diluents.
Diluent
Ingredients (g/L) BPSE Lake’s Phosphate NaCl-TES
Glucose 5.00 10.00 6.00
Magnesium chloride (6H20) 0.34 0.68
Tripotassium citrate (H20) 0.64 1.28
Sodium acetate (3H20) 4.30 8.51
Sodium glutamate (H20) 8.67 19.20
TES 1.95 1.37
Potassium monophosphate
(anhydrous) 0.75 14.56
Dipotassium hydrogen phosphate
(3H20) 12.70 8.37
Sodium chloride 8.00
Sodium hydroxide (1 M) 2.75 ml
Osmolarity (mOsmol/kg H2O) 370 368 300 382
pH 7.4 7.1 7.1 7.4
1. The number of live spermatozoa
In general, the survival of spermatozoa depended on the diluent, storage time, dilution rate and
temperature (P < 0.001). The highest survival was found with the Lake’s and NaCl-TES diluents
whereas the lowest was seen in BPSE and Phosphate diluents (Fig. 2).
0
20
40
60
80
100
Undiluted
Phosphate
NaCl-TES
Lake’s
BPSE
03624
Time (hours)
% Live spermatozoa
Figure 2. Effect of diluent and storage time on the number of live spermatozoa.
The survival of undiluted spermatozoa was equally as high in Lake’s and NaCl-TES diluents. There
was a lower number of live spermatozoa after 24 hours of storage. The survival of spermatozoa was
affected by temperature with more (P < 0.01) spermatozoa surviving at 4 than at 20°C (Fig. 3). The
least (P < 0.01) spermatozoa survived storage at 39°C. More spermatozoa (P < 0.01) survived at
dilutions 1:2, than at dilutions 1:4 and 1:8. The lowest (P < 0.01) number of spermatozoa was found
live when diluted 1:8.
15
0
20
40
60
80
100
Undiluted
Phosphate
NaCl-TES
Lake’s
BPSE
42039
Temperature (°C)
% Live spermatozoa
Figure 3. Effect of diluent and temperature on the number of live emu spermatozoa.
Survival of spermatozoa at 4°C
More than 80% of spermatozoa was found live after 24 hours of storage in every diluent (Fig. 4). After
3 hours of storage, the number of live spermatozoa did not change, but after 6 hours, the number
remained unchanged only in Lake’s diluent. At this temperature, more spermatozoa survived at
dilution 1:2, than at 1:4 and 1:8 dilutions during the storage period (P < 0.001).
Time (hours)
03624
0
20
40
60
80
100
Undiluted
Phosphate
NaCl-TES
Lake’s
BPSE
% Live spermatozoa
Figure 4. Effect of diluent and storage time on the number of live emu spermatozoa stored at
4°C.
Survival of spermatozoa at 20°C
The number of live spermatozoa stored at 20°C for 24 hours depended on the diluent. More than 90%
of spermatozoa survived storage for 6 hours in BPSE, Lake’s and NaCl-TES diluents (Fig. 5).
However, after 24 hours of storage, the number of live spermatozoa in every diluent was lower (P <
16
0.01). Undiluted spermatozoa remained unchanged. Phosphate and BPSE diluents were the least
effective in maintaining the number of live spermatozoa at this temperature.
0
20
40
60
80
100
03624
Time (hours)
Undiluted
Phosphate
NaCl-TES
Lake’s
BPSE
% Live spermatozoa
Figure 5. Effect of diluent and storage time on the number of live spermatozoa stored at 20°C.
Survival of spermatozoa at 39°C
Storage of spermatozoa at this temperature was the most detrimental and few survived 24 hours
storage in any diluent (Fig. 6). The number of live spermatozoa did not change for up to 6 hours in
Lake’s and NaCl-TES diluents, but was markedly lower in BPSE and Phosphate diluents. The number
of live spermatozoa stored in Phosphate diluent was reduced after 3 hours (P < 0.01) and it declined
further after 6 hours. The number of spermatozoa in undiluted semen was higher (P < 0.01) than in
Phosphate and BPSE after 6 hours of storage.
03624
0
20
40
60
80
100
Phosphate
NaCl-TES
Undiluted
Lake’s
BPSE
% Live spermatozoa
Time (hours)
Figure 6. Effect of diluent and storage time on the number of live spermatozoa stored at 39°C.
2. Effect of storage on motility of emu spermatozoa
17
The motility of emu spermatozoa was significantly affected by diluent, temperature, dilution and
storage time. The highest motility was maintained in NaCl-TES, Lake’s diluents and in undiluted
semen in every temperature while lower motility was scored in BPSE and Phosphate diluents (Fig. 7).
0
1
2
3
4
5
Temperature (°C)
Motility
42039
Phosphate
NaCl-TES
Undiluted
Lake’s
BPSE
Figure 7. Effect of diluent and temperature on motility of emu spermatozoa.
Increasing the storage time decreased motility. Only spermatozoa diluted in Lake’s and NaCl-TES
diluents, as well as undiluted spermatozoa, appeared to lose their motility slower than those diluted in
Phosphate and BPSE (Fig. 8).
24 h
6 h
3 h
0 h
0
1
2
3
4
5
Motility
BPSE Lake NaCl-TES Phosphate Undiluted
Diluent
Figure 8. Effect of diluent and storage time on motility of emu spermatozoa.
The motility of emu spermatozoa was affected by dilution in some diluents. The higher the dilution,
the lower the motility (Fig. 9). Spermatozoa diluted in NaCl-TES maintained their motility regardless
of the dilution rate. There was severe loss of motility with Phosphate and BPSE and intermediate loss
with Lake’s.
18
248
0
1
2
3
4
5
Undiluted
Phosphate
NaCl-TES
Lake’s
BPSE
Dilution
Motility
Figure 9. Effect of dilution and diluent on motility of emu spermatozoa.
In conclusion, the most suitable diluent for storing emu spermatozoa is probably going to be Lake’s or
NaCl-TES. For up to 3-6 hours, emu spermatozoa could be maintained in these diluents at 20°C
without the loss of viability. However, if storage for up to 24 hours is required, emu spermatozoa
would need to be stored at 4°C.
3. Effect of Na and K concentrations on the viability of emu spermatozoa at 20°C
Lake’s and the NaCl-TES diluents appear best able to preserve the viability of emu spermatozoa, but
these results did not appear to differ from the viability of undiluted spermatozoa. Using our
knowledge of the concentrations of major inorganic ions in emu seminal plasma, we modified Lake’s
diluent to formulate new diluents (UWA-E1, UWA-E2 and UWA-E3) with varying concentrations of
Na and K (Table 8).
Table 8. Characteristics of diluents that were investigated, in relation to emu seminal plasma.
Diluent
Sodium
(mM/L) Potassium (mM/L) pH mOsmol
UWA-E1
UWA-E2
UWA-E3
Lake’s
NaCl-TES
Seminal plasma
118.2
128.9
100.1
165.2
135.0
130.0
15.8
13.9
17.9
11.8
–
15.0
7.3
7.3
7.3
7.1
7.4
7.4
291
305
264
368
382
300
The effect of these diluents on the viability of emu spermatozoa at 20°C was then investigated in two
trials, but the storage time was extended to 48 hours since the loss of viability in Lake’s and NaCl-
TES appeared small after only 24 hours of storage while undiluted spermatozoa were not affected. In
the first trial, the number of live spermatozoa was determined, and in the second motility was scored.
19
After 6 hours, the number of live spermatozoa was reduced only in NaCl-TES diluent (P < 0.05).
After 48 hours, there were more than double the number of live spermatozoa in UWA-E1, UWA-E2
and UWA-E3 diluents than in undiluted semen, or in Lake’s or NaCl-TES diluents.
3 6 24 48
0
20
40
60
80
100
Undiluted
NaCl-TES
Lake’s
UWA-E3
UWA-E2
UWA-E1
Storage time (hours)
% Live spermatozoa
Figure 10. Effect of diluents on the number of live spermatozoa stored at 20°C.
The motility of spermatozoa in UWA-E1, UWA-E2 and UWA-E3 diluents was lower (P < 0.01) than
in Lake’s, NaCl-TES and undiluted semen after 1 hour of storage, and was further reduced (P < 0.01)
after 3 hours, but then remained unchanged for the rest of the storage period. The motility of
spermatozoa in Lake’s, NaCl-TES and undiluted semen declined gradually after 3 hours.
Storage time (hours)
0
1
2
3
4
5
Motility
13 6 24
48
Undiluted
NaCl-TES
Lake’s
UWA-E3
UWA-E2
UWA-E1
Figure 11. Effect of diluents on the motility of emu spermatozoa stored at 20°C.
In conclusion, our results suggest that by reducing the motility of emu spermatozoa, their viability can
be extended. It is very encouraging to find that we can extend the viability for up to 48 hours at 20°C.
The fertilising ability of spermatozoa stored in ‘UWA-E’ diluents and the mechanism responsible for
reduction of motility need to be further investigated.
Experiment 5: Development of insemination technique (AI)
In principle, any insemination technique needs to be quick and highly efficient in order to minimise
the loss of spermatozoa. Females were selected on the basis of their crouching behaviour and their
response to massage stimulation. The crouching behaviour probably develops due to imprinting to
20
humans. Not every female imprints, but if this approach could be used, the insemination technique
would be stress free, and therefore would not interfere with ovulation. Female emus can be
inseminated in two ways. We investigated the use of the speculum and the non-speculum technique
and different insemination straws.
21
AI with speculum
When the female is in the crouching position, the inseminator kneels behind her and gently inserts the
speculum into the cloaca and brings it close to the vagina, using the light from the bulb at the tip of the
speculum to see the vaginal opening. An insemination straw mounted on a tuberculin syringe is then
introduced directly into the vagina and the semen is injected into it. There are, however, some
problems associated with this technique. The speculum needs to be warmed up to a body temperature
as the female can terminate crouching on feeling a cold speculum. Secondly, not every female can
raise her tail high enough and therefore expose the vent adequately for the speculum to be inserted
easily and for the inseminator to be able to see the inside of the cloaca. This depends on two factors,
the receptivity of the female and the morphology of the tail. Certain female emus, even though they
crouch, do not show the ‘sexual excitement’ that they would normally show when mounted by a male,
so they keep their tail low. Also, some female emus have a tail morphology in which the vent is almost
horizontal with the tip of the tail, and therefore very difficult to access. This would a problem for any
insemination technique and such females would need to be excluded from AI programs.
Non-speculum AI
At first, we started with the palpation of the vagina. Briefly, the inseminator locates the vagina with
the index finger of one hand. The straw, mounted on a tuberculin syringe held in the other hand, is
then introduced into the cloaca along the index finger and inserted into the vagina. The index finger is
withdrawn and the straw is inserted further until resistance is felt. A dose of semen is introduced into
the vagina as the inseminating straw is gradually withdrawn. We found that the palpation of the vagina
can be avoided and the straw directly inserted into the vagina. This is because, when the female is
receptive and in a crouching position, the opening to the vagina is brought close to the vent. Upon
additional massage stimulation, the opening can be seen right at the vent, while the vent is slightly
open. When the insemination straw is placed into the opening and the female is still stimulated by
massaging her back and sides the vaginal opening remains at the vent for a short time that is
nevertheless sufficient to insert a straw deep into the vagina and inject semen. It is important during
this procedure that the inseminator does not place his fingers into the cloaca as this can, but not
always, cause the vaginal opening to be drawn back and deep into the cloaca and terminate the
receptivity of the female. This technique requires practise but is probably the most efficient as it feels
like the straw is being drawn into the oviduct.
In collaboration with Cook Australia Pty Ltd, several types of insemination straws were examined for
the length, volume of contents, shape of the tip and rigidity. Recommendations were sent back to the
manufacturer so that the right type of insemination straw would be made. The straws that were made
upon request did not have required rigidity, were too soft and could not be improved by the
manufacturer. We therefore had to use our own plastic straws for AI experiments.
Experiment 6: Effect of storing semen in vitro on viability of spermatozoa in vivo: Insemination
trial
This experiment could not be conducted as the development of the emu semen diluent could not be
finished and not enough females were available.
Experiment 7: Effect of semen dose on the length of the female fertile period
This experiment was designed to determine an optimal dose of spermatozoa for AI. Females (4 per
semen dose) were inseminated with 120, 200, or 400 million sperm. Semen was collected from 4
males, pooled and the concentration determined by spectrophotometry. Semen was then diluted 1:1
with Lake’s diluent and females were inseminated within 30 min of semen collection. Following a
single insemination, eggs were collected and incubated. The eggs were candled to determine fertility
during incubation at Day 7 and at Day 14 of incubation, and then they were broken to confirm
embryonic development.
22
23
The females inseminated with 120 million sperm had the same median duration of fertility as those
inseminated with 200 million sperm, while 400 million sperm produced 14 days fertility (Table 9).
Dose (million sperm) Days fertile
120 10.5
200 10.5
Table 9. Median duration of
fertility following a single
insemination with fresh semen. 400 14
Given that 120 million sperm can maintain female fertility for up to 10 days, and that an average
ejaculate contains about 2 billion spermatozoa, then 16 AI doses could be made from each ejaculate.
As semen can be collected twice a day, then 32 females inseminated by one male every 10 days. Over
one breeding season, one elite male could serve up to 320 females.
Experiment 8: Effect of the frequency of insemination on egg fertility
This experiment was carried out following Experiment 7. Five females were inseminated 3 times, once
every 10 days, with 120 million sperm. The eggs were collected and then incubated. Unfortunately,
the results of this experiment were lost when a malfunction in the incubator on 13 August 1999 ejected
most of the eggs onto the floor.
Experiment 9: Effect of twice daily collection on the maintenance of high semen yields
This experiment had to be cancelled for two reasons. First, the experiment on the effect of season on
semen quality and quantity had to be continued until mid-July 99, after which it was too late to begin
another experiment that would have to last for about 4 weeks. Secondly, in the absence of technical
assistance, it was not feasible because of the time needed to carry out the other experiments in the
program.
Experiment 10: Long term preservation — development of cryopreservation techniques
This experiment was carried out to determine the best cryoprotectants and their optimal concentration,
and cooling/freezing and thawing rates. Semen was diluted with diluent and cryoprotectant and
subjected to various cooling/freezing rates before being plunged into liquid nitrogen. The viability of
spermatozoa before and after freezing was examined by scoring motility and counting the number of
live spermatozoa following staining with nigrosine-eosin stains under light microscopy. Because
cryoprotectants are known to be toxic at certain levels, the effect of every cryoprotectant on emu
spermatozoa was first investigated.
Effect of cryoprotectant on viability of emu spermatozoa before freezing
Emu semen was collected, pooled, diluted 1:1 with cryopreservation diluent (Table 10) and semen
temperature was reduced to 5°C by holding the sample in the –20°C freezer. The semen was then
aliquoted into glass tubes held in the iced water bath and cryoprotectant was added in bulk to make a
final concentration of 3, 6, 9 and 12% of cryoprotectant in diluted semen. Three cryoprotectants were
investigated: glycerol (GLY), dimethylsuphoxide (DMSO) and dimethylacetamide (DMA).
Table 10. Composition of cryopreservation diluent (Wishart, 1995)
Chemical g/100 mL of DI H2O
24
Sodium glutamate (H2O)
Magnesium acetate (4H2O)
Potassium acetate
Polivinyl pyrrolidone (MW 10,000)
Glucose
1.92
0.08
0.50
0.30
0.80
Only glycerol appeared to affect the integrity of spermatozoal membrane, causing more than 90% of
sperm to become pink, whereas the reverse was seen for spermatozoa treated with DMA and DMSO
(Fig. 12).
% of white sperm
SEMEN
GLY
DMSO
DMA
% of pink sperm
0
60
20
40
80
100
0
20
40
60
80
100
036912
0
4
8
12
16
% of dual sperm
Cryoprotectant concentration (%)
Figure 12. Effect of cryoprotectant on the integrity of spermatozoal membrane
With DMA and GLY, the number of dual spermatozoa decreased with increased concentration. The
number of normal spermatozoa was not affected by cryoprotectant or concentration. During
morphological examination, spermatozoa treated with GLY appeared swollen whereas those treated
with DMA or DMSO appeared shrunk or normal. These results suggest that GLY readily penetrated
the spermatozoal membrane and this process allowed the dye to enter the cells. On the other hand,
DMSO and DMA would not penetrate the membrane, but due to high osmolarity of the medium
(Table 11), sperm cells lost some intracellular water and shrank.
25
Table 11. Osmotic conditions at different concentrations of cryoprotectant
Cryoprotectant Concentration (%) Osmolarity (mOsmol/kg
H2O)
GLY 3 756
6 1285
9 1798
12 2340
DMA 3 706
6 1143
9 1576
12 2100
DMSO 3 771
6 1293
9 1908
12 2560
Semen + diluent 1:1 342
Effect of cryoprotectant and freezing on the viability of emu spermatozoa
Semen was collected, pooled and diluted 1:1 with cryopreservation diluent and cooled to 5°C. The
precooled cryoprotectants were added to semen to a final concentration of 3, 6 and 9%.
Semen was held in glass tubes in iced water bath for 30 min to equilibrate and then semen was loaded
into the freezing straws and sealed. The following protocols were tested:
A. Protocol 1
Step 1. Hold at +5°C
Step 2. Reduce temperature from +5°C to –35°C at the rate of 7°C/min
Step 3. Reduce temperature from –35°C to –140°C at the rate of 20°C/min
Step 4. Put into liquid nitrogen
B. Protocol 2
Step 1. Hold at +5°C
Step 2. Reduce temperature from +5°C to –35°C at the rate of 3°C/min
Step 3. Hold at –35°C for 5 min
Step 4. Put into liquid nitrogen
C. Protocol 3
Step 1. Hold at +5°C
Step 2. Reduce temperature from +5°C to –35°C at the rate of 1°C/min
Step 3. Hold at –35°C for 5 min
Step 4. Put into liquid nitrogen
D. Protocol 4 (rapid freeze)
Step 1. Hold at +5°C
Step 2. Put into liquid nitrogen
The protocols 1, 2 and 3 were carried out using the programmable freezer.
26
Following freezing and holding for 2 days in the liquid nitrogen, the samples were then thawed in the
iced water bath (5°C), the straws cut open and semen released onto the glass slides and the number of
live spermatozoa and motility were examined.
Following rapid freezing there was no motile spermatozoa at any concentration. Therefore only
Protocols 1,2 and 3 were analysed further. There were no motile spermatozoa in the absence of
cryoprotectant, irrespective of the protocol (Fig. 13). Motility increased with increased concentration
of cryoprotectant. Increasing the concentration of cryoprotectant to 9% yielded the highest number of
live spermatozoa (Fig. 13).
Concentration of cryoprotectant (%)
0369
0
10
20
30
40
0
1
2
3
0369
Motility
% Live sperm
Figure 13. Effect of cryoprotectant concentration on motility and number of live emu spermatozoa
following freezing.
The slower the freezing rate the higher the number of live spermatozoa. DMA yielded more live
spermatozoa than DMSO and GLY (Fig. 14).
In conclusion, freezing at the rate of 1°C/min with 9% of DMA appears to be optimal for emu
spermatozoa.
Concentration of cryoprotectant (%)
0369
0
10
20
30
40
0
1
2
3
0369
Motility
% Live sperm
27
0
10
20
30
40
50
3
2
1
% of live sperm
36 9
DMA
369
DMSO
369
GLY
Protocol
Figure 14. Effect of freezing protocol and cryoprotectant on the number of live emu spermatozoa.
28
5. Discussion
The production of semen and spermatozoa is very seasonal, lasts for about 6 months and can be
divided into 3 periods: the beginning, middle and the end. It appears that, at the beginning of sperm
production, it takes more time for males to attain the peak of production than it takes to decline from
the peak to its termination. Semen can be collected as early as March, which is about when female
emus start egg laying, but the libido of males is not as high as in the middle of the breeding season and
initially semen can only be collected once a day. The output of spermatozoa is also lower than for the
rest of the season as the concentration of semen is low. On the other hand, the quality of spermatozoa
appears to be better. Ejaculates contain more live spermatozoa and there is more unsaturated fatty
acids in spermatozoa. Due to the lower output of spermatozoa, from one ejaculate only about 8 AI
doses could be made. A demand for spermatozoa to inseminate females early in the season could be
met by selecting males that start semen production early in the season. On the other hand, the lower
output of spermatozoa might be offset by their greater retention in the female oviduct. Better retention
of spermatozoa at the start of egg laying has been reported for the chicken and turkey (Brillard, 1994),
so the female emus could be retaining more spermatozoa at the beginning of egg laying. This suggests
that less spermatozoa might be needed to maintain 10 day fertility, or perhaps the females could be
inseminated at a greater than 10 day interval. These requires further studies.
As the production of semen and spermatozoa reaches its peak in June-July, there is more spermatozoa
available for AI and about 23 females could be sired from one ejaculate. As semen can be collected
twice a day (Malecki et al. 1997b), twice as many females could be inseminated. Nevertheless, the
production of spermatozoa and their quality soon starts decreasing and the number of females that can
be inseminated from one ejaculate reduces by about 30%. This time of the breeding may be
additionally affected by females having lower ability to retain spermatozoa and also by some males
stopping their semen production. The female factor is not known yet, however, but it has been
demonstrated in the chicken and turkey (Brillard, 1994) so we can anticipate it in the emu. Studies are
now needed to evaluate it.
It needs to be stressed that the results of this study come from a selected group of birds maintained at
the research facility. These birds, as with those on commercial farms, have not been subjected to much
selection pressure and, as the variation between individuals suggests, there is a lot of room for
selection in a number of traits, such as semen output, libido, egg production and fertility. Once we
have introduced proper selection programs on farms, the efficiency of production should increase and
this would further benefit the producers.
Good storage and preservation techniques are essential in any AI program. The emu semen diluent
needs development but the results presented here are very encouraging. Because the levels of
potassium, sodium, calcium and magnesium in emu seminal plasma were similar to those in the
chicken and turkey, we anticipated that commercial poultry diluents should be suitable for storing emu
semen. Our studies clearly show that only Lake’s diluent can be recommended. Emu semen can be
stored in Lake’s diluent at either 4 or 20°C for up to 6 hours without the loss of viability. However,
these results need to be verified by studies of the fertilising ability of spermatozoa because, in our
experiments, we only evaluated the integrity of spermatozoal membrane and spermatozoal motility,
parameters that are not highly correlated with fertility (Wishart, 1984).
Storage in the newly developed emu diluents should result in more live spermatozoa. Our diluents
(UWA-E1, UWA-E2 and UWA-E3) maintained more viable spermatozoa than any other diluent and
our results indicate that sperm viability can be extended by reducing motility and that this could be
achieved by increasing the concentration of potassium and reducing sodium in the diluent. Again, we
still need to test the fertilising ability of spermatozoa following storage in such diluents.
Emu spermatozoa can be frozen in liquid nitrogen. They have low susceptibility to cold shock and a
slow cooling/freezing rate is most efficient. The recovery of viable spermatozoa (40%) is as good as in
the chicken, but we should be able to improve it further because our freezing trials were carried from
the end of July when the quality of spermatozoa begins to decrease. We expect that spermatozoa
29
frozen at the beginning and middle of season would have better freezing potential. In addition, we
only used pooled semen. In poultry, there are differences between males in the ability of their sperm to
survive the freezing/thawing process (Hammerstedt, 1995) and we expect similar differences between
individual male emus. We have not yet tested this. The difference between individuals in sperm
freezing potential may be due to the fatty acid content of their spermatozoa, which depends on
nutrition and age (Kelso, 1997), as well as season (this study). This is another area where we could
improve the freezing potential of emu spermatozoa.
Deposition of spermatozoa in the female oviduct (AI) can be successful and efficient if there is good
cooperation between the female and the inseminator. The crouching behaviour that female emus
develop allows for a stress free approach, and therefore would not interfere with ovulation. The
insemination technique can be learned and used, and little intervention in the cloaca is needed because
the non-speculum technique has also proven to have potential.
An elite male producing average ejaculates could be used to fertilise 32 females every 10 days, or up
to 320 females in a season. We know, however, that the output of spermatozoa is seasonal and the
male:female ratio will vary depending on the period of breeding season. Nevertheless, the
considerable variation in sperm production between males (Malecki et al. 1997b) means that high
sperm producers could be selected and the male:female ratio could be increased further. Artificial
insemination could also be made more efficient by storing semen in the right diluent and by a higher
dilution of semen before AI which reduces viscosity. In addition, the efficiency of AI could be
improved by selecting females that show a longer duration of fertility or a better retention of
spermatozoa in the oviduct.
In conclusion, the production of spermatozoa, even though it is affected by season, could be very
effectively used in artificial breeding programs for the emu. Emu farming can already benefit from the
current techniques for semen collection, sperm storage, preservation and artificial insemination, and
thus begin selection programs. Much of the genetic potential of this bird is still to be explored.
30
6. Implications
We are still not certain of the fertilising ability of emu spermatozoa following storage and
cryopreservation because the conventional techniques (sperm cell integrity and motility), that we used,
overestimate the fertilising ability of stored or cryopreserved spermatozoa. The fertility trials would
most accurately estimate this functional ability, however, a new approach of using the sperm-egg
interaction assay could be developed, and this would not require to carry out the expensive, time-
consuming AI trials. that need large numbers of females, daily egg collection and time-consuming
incubation. The sperm-egg interaction assay could have wide application from fertility assessment of
individual birds to the entire flocks. From this technology would benefit both, the emu and ostrich
producers.
By introducing reproductive technology, the emu industry could now make use of the massive
reservoir of elite genes and genetic diversity of the emus. Our farmed flocks could be improved by
selection and introduction of new genes.
In addition, artificial insemination can substitute for natural mating and greatly reduce the male to
female ratio, saving on feed costs and capital resources, which can then be used to manage more
females and therefore produce more eggs and growing chicks.
31
7. Recommendations
We have demonstrated that emus can be bred by artificial insemination. Good quality ejaculates can be
collected from males into the artificial cloaca by two methods. Using these methods, the male emus
could be trained and their sperm production potential assessed. Similarly, female emus can be selected
for the breeding program based on their egg laying records and their crouching behaviour. The emu
diluent is yet to be fully developed, but spermatozoa for artificial insemination can be stored
successfully for up to 6 hours at 4 or 20°C in Lake’s diluent diluted 1:2.
Finally, emu spermatozoa can be cryopreserved for sperm banking or for transport over long distances.
At present, the best recovery is achieved by freezing pre-cooled (to 5°C) sperm samples containing 9%
DMA at the rate of 1°C/min to –35°C, and then plunging them into liquid nitrogen.
As season will affect the availability of spermatozoa for AI, early and late sperm producing males
could be sought to meet these demands. Males that produce sperm from March until September would
be best, and we might be able to select them from existing flocks, or breed selectively for longer
duration of sperm production. This is one of many breeding objectives that emu farming can now
focus on with this technology.
32
8. Bibliography
Allen C. J. and Champion L. R. (1955), Competitive fertilization in the fowl. Poultry Sci., 34, 1332-
1342.
Chalah, T. and Brillard, J. P., 1998. Comparison of assessment of fowl sperm viability by eosin-
nigrosin and dual fluorescence (SYBR-14/PI); Theriogenology, 50: 487-493.
Chaudhuri, D. and Lake, P. E. 1988. A new diluent and method of holding fowl semen for up to 17
hours at high temperature. 18th World Poultry Congress, Nagoya, 591-593.
Brillard, J. P., 1994. Artificial insemination: How many sperm, How often? Pages 176-183 in
Proceedings of the 1st International Symposium on the Artificial Insemination of Poultry, 17-19 June
1994, University of Maryland, College Park, USA.
Donoghue, A. M., Garner, D. L., Donoghue, D. J., and Johnson, L. A. 1995, Viability assessment
of turkey sperm using fluorescent staining and flow cytometry. Poultry Sci 74: 1191-1200
Etches, R., 1996. Artificial insemination. Pages 234-262, in Reproduction in Poultry, CAB
International, UK.
Hammerstedt R. H., 1995. Cryopreservation of poultry semen - current status and economics, First
International Symposium on the Artificial Insemination of Poultry., pp 229-250, Eds; M. R. B. a. G. J.
Wishart. Poultry Science Association, University of Maryland, College Park, USA.
Kelso K. A., Cerolini S., Speake B., Cavalchini L. G. and Noble R. C., 1997. Effects of dietary
supplementation with a-linoleic acid on the phospholipid fatty acid composition and quality of
spermatozoa in cockerel from 24 to 72 weeks of age. J. Reprod. Fert., 110, 53-59.
Lake, P. E., 1960. Studies on the dilution and storage of fowl semen. J. Reprod. Fert. 1: 30-35.
Lake, P. E., 1986. The history and future of the cryopreservation of avian germ plasm. Poultry Sci.
65: 1-15.
Lake, P. E. and Ravie, O. 1984. An exploration of cryoprotective compounds for fowl spermatozoa.
Brit. Poultry Sci. 25: 145-150.
Lake, P. E. and Wishart, G. J. 1984. Comparative physiology of turkey and fowl semen, Pages 151-
160 in: Reproductive Biology of Poultry (eds. F.J. Cunningham, P. E. Lake & D. Hewitt) British
Poultry Science Ltd., London, UK.
Lake, P. E. and J. M. Stewart, 1978a. Artificial Insemination in Poultry, Bulletin 213, Ministry of
Agriculture, Fisheries and Food, London, Her Majesty's Stationery Office.
Lake, P.E. and Stewart, J. M. 1978b. Preservation of fowl semen in liquid nitrogen-an improved
method. Brit. Poultry Sci. 19: 187-194.
Malecki I. A., Martin G. B. and Lindsay D. R. (1997a), Semen production in the emu (Dromaius
novaehollandiae). 1. Methods for collection of semen. Poultry Sci., 76, 615-621.
Malecki I. A., Martin G. B. and Lindsay D. R. (1997b), Semen production in the emu (Dromaius
novaehollandiae). 2. Effect of collection frequency on the production of semen and spermatozoa.
Poultry Sci., 76, 622-626.
Malecki I. A., Martin G. B., O'Malley P. J., Meyer G. T., Talbot R. T. and Sharp P. J. (1997),
Endocrine and testicular changes in a short-day seasonally breeding bird, the male emu (Dromaius
novaehollandiae) in the south-western Australia. Anim. Reprod. Sci., 53, 143-155.
Malecki, I. A., O'Malley, P. and Martin, G. B. 1996. Length of the fertile period in the female emu
(Dromaius novaehollandiae). The 13th International Congress on Animal Reproduction, June 30-July
4, 1996, Vol. 2, P 9-13.
Sexton, T. J., 1980. A new poultry semen extender. 5. Relationship of diluent components to
cytotoxic effects of dimethylsulfoxide on turkey spermatozoa. Poultry Sci. 59: 1142-1144.
Sexton, T. J. and Fewlass, T. A. 1978. A new poultry semen extender. 2. Effect of the diluent
components on the fertilizing capacity of chicken semen stored at 5°C. Poultry Sci. 57: 277-284.
Wishart G. J., 1984. Effects of lipid peroxide formation in fowl semen on sperm motility, ATP
content and fertilizing ability. J. Reprod. Fert., 71, 113-118.
Wishart G. J., 1995. Cryopreservation of avian spermatozoa in: Cryopreservation and Freeze-Drying
Protocols. 167-177pp. Eds; J. G. Day and M. R. McLellan. Humana Press, Totowa, NJ.
Wishart, G. J., 1989. Physiological changes in fowl and turkey spermatozoa during in vitro storage.
Brit. Poultry Sci. 30: 443-454.
33
