Exploring the relationship between larvae
abundance and individual caribou health
R.A. Been
Faculty of Veterinary Medicine
Utrecht University
July until September 2011
Abstract
It is known that gastrointestinal nematodes can have a significant impact on the growth of
farmed ruminants. The clinical signs are often subtile, but production losses can be of big
importance. Little is known about the impact of gastrointestinal nematodes in wild ruminants. I
used cross-sectional data to explore the relationship between larvae abundance and individual
caribou health. Being part of a two year master project, I evaluated results of 18 samples of
adult caribou from the Akia-Maniitsoq herd, isolating larval nematodes from the mucosal lining
of the abomasa using a pepsin-HCl digestion. The goal of this study was to discover a
relationship between mucosal larvae abundance and caribou health condition. This study did
not discover a relationship unfortunately, but these results are preliminary as this study is part of
a two year project.
Introduction
Natural and translocated populations of caribou and semidomesticated Norwegian reindeer (Rangifer tarandus spp.)
are present along the west coast of Greenland from the
southern peninsula to Inglefield Land in the north, see figure
1. My work focuses on the one of the two largest herds in
west Greenland, Akia-Maniitsoq (AM). The herd inhabits a
semi-isolated range due to the Greenlandic ice cap to the
North and extensive fjord system to the south. The Northern
people traditionally have a close relationship with caribou.
Caribou are the basis of their cultures and still play a central
role in their lives, caribou are harvested annually for meat
and outfitting (Albon et al., 2002; Melgaard, 1986; Stien et
al., 2002).
KS collection sites
KS herd range
AK collection sites
AK herd range
Austmannadalen region
Historical Norwegian
reindeer range
Figure1. Ranges of native and imported species of
interest in west Greenland
Caribou are grazers that include a wide variety of plants in their diet, but eat most lichen and
sedges. During foraging they might also ingest different kinds of gastrointestinal parasites
through fecal contamination of pastures (Albon et al, 2002; Stien et al., 2002). This could
include parasites from semi-domesticated Norwegian reindeer (Rangifer tarandus tarandus) and
domestic sheep (Ovis aries), which were imported into ranges adjacent to AM. (Melgaard,
1986).
The goal of this study was to define a relationship between larvae abundance and individual
caribou health. My work is part of a bigger two year master project. I hypothyse that caribou
health is negatively associated with larvae abundance, as it is generally accepted that gastro
intestinal parasites can have negative effects on individuals in relation to the individual’s health.
In the herd of caribou the following groups of parasites were identified through fecal analysis:
nematodes (identified by the presence of Trichostrongylid-type, Marshallagia spp. and
Nematodirine eggs) and cestodes (eggs present are most likely Moniezia spp.). The nematode
lifecycle can involve a stage of inhibition delaying maturation of ingested 3rd stage larva into
reproductive adults, occurring during unfavourable conditions for eggs on pasture, e.g. winter or
dry seasons. (Hoar et al., 2009; Smith, 1979). This delays the release of eggs until pasture
conditions favour egg survival and development, such as warmer temperatures during spring.
This resumption has effects on animal health as emerging larvae cause gross damage to the
gastric glands of the abomasums where they sequester during inhibition. Documented clinical
effects of type II Ostertagiasis, in domestic species include diarrhea, leading to a decrease in
body weight and mortality. (Albon et al, 2002; Stien et al, 2002; Hughes et al., 2009)
Procedure
Female caribou and their calves-at-heel were collected opportunistically as part of the
CircumArctic Rangifer Monitoring and Assessment (CARMA) Network initiatives. Collections
occurred from Mar. 29 to Apr. 13, 2008. Sample collections from mature animals (≤ 1 year),
including the removal and freezing of the abomasa, occurred within a few hours of caribou
being shot (Russell et al., 2010). I isolated larval nematodes from the mucosal lining of the
abomasa of 18 adult female caribou using a pepsin-HCl digestion (Appendix 1). This is a
quantitative technique, allowing us to estimate larval nematode intensity (Eysker, 1999).
Results and discussion
All of the 18 adult female caribou examined in this cross-sectional study were parasitized by
abomasal nematodes. My work only contains adult animals as I had limited time to do all
digestions and analysis of the samples needed to be able to discover a relationship between
mucosal larvae abundance and caribou health condition. Testing for negative effects of
nematodes on caribou health condition in this study is difficult because both small sample size
and the variation among samples (animal age, body mass, body condition) can add to the error
variance. (Gunn, 2008; Stien et al. 2002)
Because in this study there were only adult samples analyzed, there are several points in time
missing, for example the time span wherein the calves immune response develops is not
covered in this study. This research is part of a bigger Masters project at the University of
Calgary (by Jillian Steele), the results that were available at the time this paper was written are
only preliminary results for the bigger project that will also cover other age classes of caribou.
The final research will contain the results of caribou from two adjacent herds, Akia-Maniitsoq
(AM) and Kangerlussuaq-Sissimiut (KS). In total there will be results from 40 AM samples and
33 samples from KS. I will discuss my results out of the 18 adult samples in comparison with
previous work that was done, there is no statistical analysis of the results included as my data
have shown no statistical significance, most likely due to the small sample size. (Appendix II)
Nematode demographics
Nematode demographics were estimated by analyzing the abomasal digests as described in the
procedure. The abomasa were collected during post-mortem examination in the spring of 2008.
Results estimated from abomasal digests
The results of the abomasal digests showed that the larva abundance varies between 265 and
2000 larvae per caribou. It also shows that intensity of infection with larvae in the abomasal
mucosa rises when the host age increases. There was one animal that differed a lot from the
others; this animal had a mucosal larvae count of 2000 larvae.
Previous research has confirmed that nematode abundance increases with host age, reaching
asymptotic levels at 2-4 years of age for O. gruehneri and at about 4 months of age for M.
marshalli (Halvorsen et al.,1999; Irvine et al,. 2000). The results found in this study showed no
statistical correlation for nematode abundance and caribou age, but this might be due to the
small sample size and the small variation in the adult age class.
Relation to body condition
In wild ungulates fat reserves are widely accepted to be a good reflection of general body
condition, and we assessed possible relations between these, general body parameters (e.g. body
weight), reproductive status and larval nematode abundance. (Allaye Chan-McLeod et al.,
1995)
Body weight
Previous work has shown a significant decrease in body weight with increasing abundance of
mucosal larvae, and this effect was exaggerated in non-pregnant animals of the population.
(Halvorsen & Bye 1986, 1999). In this study I found no correlation between larva abundance,
body weight and pregnancy, but as body weights are highly variable with respect to age and
season of the year, we likely need to add more parameters to the assessment.
Back fat
Back fat depth is considered a reliable indicator of ungulate health condition, it is generally
considered better than weight since weight depends on animal size as well (Gerhart et al., 1996).
Moreover, Adamczewski et al. (1987) found that the relation between total dissectible fat and
depth of back fat is linear or near-linear in Rangifer, this shows that the use of back fat is a good
index for total body fat. In this research no relation was found between measures of back fat
depth and larva abundance. This is similar to results found by Huges at al., 2009 and Stien at
al., 2002. Back fat is the first fat reserve to be used during malnutrition; therefore there is no
correlation below 6% total body fat, which as all our animals are in poor body condition, may
be the case.
Riney kidney index
Visceral fat is deposited around the internal organs, such as kidneys, in ungulates when food
availability is good. The Riney kidney index (RKI) is proven to have good correlation with
dissectible fat, moreover at the low range of body condition it is expected to be a good index.
The use of the RKI which compares kidney and kidney fat weight, has been criticized as kidney
weights may fluctuate during the year, by Batcheler and Clarke 1970; Dauphine 1975. Our
samples were collected during spring 2008, all samples were collected during the spring so the
fluctuation of kidney weight during the year should not be of importance, since all samples were
collected in the same time of year. We found no correlation between larva abundance and
kidney fat, which is surprising as the RKI works well in animals with low fat reserves, making it
preferred over back fat depth (Riney, 1955).
Kidney fat is more mobile than marrow fat, so when condition of the caribou declines, kidney
fat will deplete first, before other fat depositions will be used (Kie, 1988). This points out that
when an animal has no kidney fat, it might still be in a condition to survive and reproduce. It is
important to realize that it will probably have other effects on the animals health condition,
those effects will not be discussed here.
Riney kidney index values can range from a low of only a few percent, representing solely
connective tissue and no fat, to over 100 percent. The animals in this study were all in a range
between 18 and 136 % RKI, with an average of 54%.
Bone marrow fat
Bone marrow fat is known to have a good correlation with body fat in thin, starving animals.
Since all our animals are in poor condition, one might expect to find a relationship between
bone marrow fat and larva abundance. In this study I did not find a relationship between bone
marrow fat and larva abundance, which could also be a result of the small sample size. (Kie,
1988; Neiland 1970).
Future research
Testing for relations between nematodes and caribou health in this study is challenging because
of both the small sample size and the variation among samples (animal age, body mass, body
condition) which can add to the error variance in small sample sizes. It is also complicated by
our lack of knowledge of nematode abundances and host health over a period of time, as body
condition is not affected by current nematode abundance, but rather the burden of past times
(Stien et al., 2002).
There are several other reasons for this cross sectional study not detecting parasite impacts on
caribou body condition. First, both negative and positive relationships are possible. For
example, body mass may be negatively related to worm burden but fitter individuals such as
those that have become pregnant may have higher worm burdens. Second, individuals that eat
more may have more parasites and therefore may be heavier, in better condition and better
able to withstand high parasite loads. Some of these problems can be overcome by conducting
experiments that manipulate parasite loads which allows a direct test of the impact of parasites
on host body condition scores (Côte et al., 2005; Hutchings, 2001; Stien et al., 2002).
As my work contains only adult animals, future work is needed with a larger sample size and
more complex models covering other age classes of caribou; sub-adults and calves. This would
let us examine the time span wherein the immune response develops which is not covered in
this study. Other considerations regarding our results include other factors influencing the
health of the animal as well, for example the time of year, animal diseases and availability of
food. Second is that in this study there was no differentiation between the different nematode
species, previous work has shown that the different species vary in the time of year when their
abundance peaks.
Acknowledgements
I want to thank Deborah van Doorn from Utrecht University for her assistance from the
Netherlands and for giving me the opportunity to fulfill my research project on the subject of
parasitology.
I want to thank dr. Susan Kutz, dr. Karin Orsel and Jillian Steele from the University of Calgary
for the opportunity to do my research project at their university. James Wang for always helping
me out with troubles in the lab, Robbert Huggins for his help in the statistical part of this
research, Marianne Vervest and Cynthia Kashivakura for the fun times in the lab and last but
not least Barb Cowley for her great hospitality, thanks to them all I had a very nice time working
and living in Calgary.
References
Adamczewski, J. Body composition in relation to seasonal forage quality in caribou (Rangifer tarandus
groen-landicus) on Coats Island, Northwest Territories. MSc., 1987 U. Alberta
Albon, S.D., Stien, A., Irvine, R.J., Langvatn, R., Ropstad, E. & Halvorsen, O. The role of parasites in t
he dynamics of a reindeer population. Proceedings of the Royal Society Biological Sciences Series
B, 2002, 269(1500): 1625-1632.
Allaye Chan-McLeod, A.C., White, R.G., Russel, D.E. Body mass and composition indices for female
Barren-Ground caribou. The Journal of Wildlife Management, 1995, 59(2), p.278-291)
Côte, S.D., Stien, A., Irvine, R.J., Dallas, J.F., Marshall, F., Halvorsen, O., Langvatn, R., Albon, S.D.
Resistance to abomasal nematodes and individual genetic variability in reindeer. Molecular
Ecology, 2005, 14(13), p. 41594168
Eysker, M., Klei, T.R., Mucosal larval recovery techniques of cyathostomes: can they be standardized?
Veterinary Parasitology, 1999. 85, p. 137–149
Gerhart, K.L., White, R. G., Cameron, R. D., Russell, D. E. Estimating Fat Content of Caribou from
Body Condition Scores The Journal of Wildlife Management, 1996, 60 (4), p. 713-718
Gunn, A., Nixon, W., eds. Monitoring Protocols - Level 2. Rangifer health and body condition
monitoring, ed. CARMA. 2008, p. 54.
Halvorsen, O., Stien, A., Irvine, J., Langvatn, R. & Albon, S. Evidence for continued transmission of
parasitic nematodes in reindeer during the Arctic winter. International Journal of Parasitology,
1999. 29, p. 567-579
Hoar, B., Oakley, M., Farnell, R., Kutz, S., Biodiversity and springtime patterns of egg production and
development for parasites of the Chisana Caribou herd, Yukon Territory, Canada. Rangifer 2009.
29 (1), p. 25 – 37.
Hughes, J., Albon, S.D, Irvine, R.J., Woodin, S., Is there a cost of parasites to caribou? Parasitology,
2009. 136, p. 253-265
Hutchings, M. R., Kyriazakis, I., Gordon, I. J., Herbivore Physiological State Affects Foraging Trade-Off
Decisions between Nutrient Intake and Parasite Avoidance. Ecologie, 2001. 82 (4), p. 1138-1150.
Irvine, R.J., Stien, A., Halvorsen, O., Langvatn, R., Albon, S.D. Life-history strategies and population
dynamics of abomasal nematodes in Svalbard reindeer (Rangifer tarandus platyrhynchus).
Parasitology, 2000, 120(3), p. 297-311.
Kie, J.G., Performance in Wild Ungulates: Measuring Population Density and Condition of Individuals
(1988)
Korsholm, H., Olesen, C.R., Preliminary investigations on the parasite burden and distribution of
endoparasite species of muskox (Ovibos moschatus) and caribou (Rangifer tarandus groenlandicus)
in West Greenland. Rangifer 1993. 13 (4), p. 185-189.
Melgaard, S., Bentounsi, B., Zouyed, I. and Cabaret, J. The Greenland Caribou – Zoogeography,
Taxonomy, and Population Dynamics, Kommissionen for Videnskabelige Undersøgelser I
Grønland, Copenhagen. 1986, 88pp.
Riney, T. Evaluating condition of free-ranging red deer (Cervus elaphus) with special reference to New
Zealand. New Zealand Journal of Science, 1955. 36, p. 429-463.
Russell, D., CARMA Network. 2010; Available from: http://www.carmanetwork.com.
Smith, H.J., Observations on the resumption of development of inhibited Ostertagia, Cooperia and
Nematodirus infections in calves stabled overwinter. Canadian Journal of Veterinary Research
1979. 43(4), p. 434–439.
Steele, J., Risk factors associated with parasite abundance and host translocation in West Greenland
caribou. AINA Grant-in-Aid proposal, 2011.
Stien, A., R.J. Irvine, E. Ropstad, O. Halvorsen, R. Langvatn, and S.D. Albon. 2002. The impact of
gastrointestinal nematodes on wild reindeer: experimental and cross-sectional studies. Journal of
Animal
Ecology 71:937.
University of Calgary, Canada, Standard Operating Procedure.
Addendum
Appendix l
University of Calgary
Department of Veterinary Medicine
Standard Operating Procedure
Procedure Title:
Method for digesting abomasa to quantify larval nematodes
Minimum Review Requirements:
Yearly
Creation Date:
30 May 2011
Date of Next Review:
30 June 2011
Administrator of Procedure:
Supervisor of Procedure:
Authorized by:
Table of Contents
Title Page and Table of Contents
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
Version History ...................................................................................................... 8
Introduction ........................................................................................................... 1
Definition ................................................................................................................ 8
Personnel ................................................................................................................ 8
Safety ...................................................................................................................... 8
Procedure ............................................................................................................... 3
Equipment or Materials Required ..................................................................... 10
Highlights / Critical Control Points ................................................................... 10
Reporting.............................................................................................................. 10
Regulatory / Standards ....................................................................................... 10
Trouble Shooting ................................................................................................. 10
References ............................................................................................................ 10
1. Version History
Version #:
Supersedes:
1.1
n/a
The signatures below indicate the person(s) responsible to administer
and supervise this procedure have read and agree toabide by the SOP
attached
Date
Handwritten amendments to the official procedures can be made by a single line through the
text, along with the date, and initialed by the authorized individual making the correction.
Changes are to be noted below. Formal changes to thisSOP are made on the date of revision or
sooner, where required.
Section
Changes made to official copy
Date
Initials
2. Introduction
This technique is used to isolatelarval nematodes from the mucosal lining of the abomasa.
This is a quantitative technique and can be used to produce estimates of larval nematode
burden and specimens for identification of species composition.
3. Definition
N/A
4. Personnel
SOP issued under authority of:
Persons authorized to perform The SOP:
Dr. S. Kutz, Jian Wang
All lab staff
5. Safety
Abomasa to beprocessed are of unknown pathogenicity (bacterial, fungal, viral and parasitic
pathogens are all possible).Proper biosafety procedures should be followed for this process.
These include:
- Always wearing PPE (gloves and a lab coat) in exposure situations
- Abomasal tissue should be disposed in a black garbage bag, double bagged
- Bags should either be taken for incineration immediately or placed in the freezer
to be incinerated at a later date
- HCL is a highly corrosive chemical and great care should be taken not to spill, imbibe or
inhale it
- If any personal exposure occurs proper first aid and biosafety procedures for
such an incident should be followed
- All waste should be disposed of in the appropriate liquid waste container
- All tools and equipment should sit in a Vircon solution (10g/L) for at least 10mins
before being cleaned with soap and water and then disinfected with ethanol
- Wash hands and forearms after performing the diagnostic test
As this procedure also involves using surgical equipment (carving knifes/ metal depressor),
care should also be taken not to cut oneself. If a wound is caused with the equipment proper
first aid and biosafety procedures for such an incident should be followed.
6.Procedure
6.1
Remove abomasa from freezer and place in a small tray; allow to defrost
6.2
Mix together pepsin-HCl solution in an Erlenmeyer (thin-necked) Flask
- Small abos (caribou) will need 1L, large abo (muskox) 1.5L
- Can be done in fume hood if technician is sensitive to HCl
- 500ml; 4g pepsin / 4.25g NaCl / 10mL HCl / Distilled H2
6.3
Place thawed abo. into the flask and seal with parafilm
6.4
Incubate the flask in a shaking incubator for 1.5hr at 39°C
6.5
Mix together another 500mL of pepsin-HCl solution
6.6
Remove flask from incubator and place the abo. on the plastic tray
6.7
Using a blunt metal depressor (or knife) scrape the mucosa off the musculature
- Should render the abo. almost see-through if done correctly
6.8
Rinse the abo., tools and your gloves with the extra pepsin-HCl solution and pour
this and the mucosalscrappings back into the flask
- Should rinse at least twice, or until volume in flask is ~1.5L
(if started with 1L)
- Abo. can now be discarded
6.9
Reseal the flask and place it back into the incubator for another 2-3hr at 39°C or
until large abo. chunks are dissolved
6.10 Remove the flask, and pour contents into a large mouthed beaker. Add distilled
water to bring volume up to a multiple of 500mL (e.g. 1L / 1.5L / 2L)
6.11 Take two 10% aliquots from the beaker
- While holding a sample cup, agitate the contents with your gloved hand
- In one go, remove 10% (e.g. 100mL / 1L; 150mL / 1.5mL)
- If excess is pulled up immediately pour it off, otherwise retake
6.12 Drain each sample through a #400 sieve and rinse to remove pepsin-HCl
- Sample must be drained into a metal hazardous liquid collector,
subsequent rinses can go into the sin
6.13 Rinse the drained sample into a clean, labeled sample container using 70%
ethanol, add enough ethanol to bring volume up to 100ml
UCID
ABOWASH
DATE
Animal ID
ETOH (70%)
1/2
6.14
6.15
6.16
6.17
Add a waterproof label (as above) and close
- Seal with parafilm if being stored before examination
All excess pepsin-HCl solution and digestion should be disposed of in the
hazardous liquid disposal
- Record amount on disposal record
The tray, beakers and tools should be washed as per lab protocols. The lab bench
should be wiped down and disinfected.
Sieve should be washed in the sonicator when procedure is complete
7.Equipment or Materials Required
Materials
7.1
Small plastic tray
7.2
1.5L Erlenmeyer flask or 2L Beaker
7.3
Parafilm
7.4
Shaking Incubator
7.5
Blunt metal depressor or dull knife
7.6
500mL Erlenmeyer flask
7.7
#400 Sieve
7.8
Specimen Cups (3)
7.9
Waterproof paper
7.10 Grease marker or pencil
7.11 Metal hazardous liquid collector
Materials
7.12 Pepsin – Should be kept refrigerated once opened
7.13 Ultrapure NaCl
7.14 HCl– Should be kept in Acid storage
7.15 70% Ethanol
8.Highlights / Critical Control Points
8.1
Concentration of pepsin-HCl solution should not be modified, nor should
temperature of incubator increased beyond 39°C
9.Reporting – To be recorded in a blue lab book
9.1
Sample notes (i.e. Date, UCID, AID, Abo. Weight, Processing Date) should all be
recorded before starting procedure
9.2
All times should be recorded in book
- First incubation start ____; Removed at _____
- Second incubation start _____; Removed at _____
9.3
Volume of final dilution and aliquots
- e.g. 1.5L ; 150mL
10.Regulatory / Standards
N/A
11.Trouble Shooting
11.1 Incubation length for the second incubation can be increased if large pieces of
abomasum are still present in digest, should always be checked at 2hr
12.References
Procedure Title:
Method for the quantification and isolation of larval nematodes from abomasal
digestions
Minimum Review Requirements:
Yearly
Creation Date:
30 May 2011
Date of Next Review:
30 June 2011
Administrator of Procedure:
Supervisor of Procedure:
Authorized by:
Table of Contents
Title Page and Table of Contents
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
Version History .................................................................................................. 8
Introduction ........................................................................................................ 1
Definition ............................................................................................................ 8
Personnel ............................................................................................................ 8
Safety................................................................................................................... 8
Procedure ............................................................................................................ 3
Equipment or Materials Required ................................................................... 10
Highlights / Critical Control Points ................................................................ 10
Reporting .......................................................................................................... 10
Regulatory / Standards .................................................................................... 10
Trouble Shooting .............................................................................................. 10
References ........................................................................................................ 10
6. Version History
Version #:
Supersedes:
1.1
n/a
The signatures below indicate the person(s) responsible to administer
and supervise this procedure have read and agree toabide by the SOP
attached
Date
Handwritten amendments to the official procedures can be made by a single line through the
text, along with the date, and initialed by the authorized individual making the correction.
Changes are to be noted below. Formal changes to this SOP are made on the date of revision or
sooner, where required.
Section
Changes made to official copy
Date
Initials
7. Introduction
This technique is used to analyse abomasal digest samples in order to quantify and isolate
larval nematodes.
8. Definition
N/A
9. Personnel
SOP issued under authority of:
Persons authorized to perform The SOP:
Dr. S. Kutz, Jian Wang
All lab staff
10. Safety
As all materials are preserved in 70% ethanol the pathogenicity of the sample is vastly
reduced, however standard laboratory safety procedures should still be followed.
- Standard PPE (Lab coat and gloves) should be worn at all times
- Wash hands and forearms after performing the analysis
6. Procedure
6.1
Take the sample to be examined and drain it through a #400 standard sieve
6.2
6.3
6.4
6.5
6.6
6.7
6.8
- Ethanol can be drained into the sink directly
With water, rinse the sample container and label into the sieve and then rinse the
sample repeatedly to remove any traces of ethanol
Rinse the contents back into the sample container with water and fill to ~100mL
Taking a 3mL bulb pipette with the tip cut off aliquot a small portion of the
rinsed sample onto a grid petri dish, add water to dilute and examine on a
dissecting microscope at 3.2.
Count larva, using a desk counter, and remove them into a small cryotube filled
halfway with 70% ethanol using a 10μL pipette
- Once having ejected a larva into the cryotube dip the pipette tip into
water so as to not agitate the next larva by adding ETOH to the petri-dish
Once full dish has been examined, rinse into a large plastic beaker and repe
When full sample has been examined place a freezer proof label on the cryotube
and place into -80°C freezer for storage
Discard waste down the laboratory sink
UCID
ABO. DIGEST (sample #)
LARVA x (# in tube)
AID
ETOH
DATE
7.Equipment or Materials Required
Materials
7.1
#400 standard sieve
7.2
Grid petri-dish
7.3
3mL bulb-pipette with the tip cut-off
7.4
Cryotube
7.5
10μL pipette and tips
7.6
Large plastic beaker for waste
7.7
Dissecting microscope
7.8
Water squirt bottle
Reagents
7.9
70% ethanol
8.Highlights / Critical Control Points
N/A
9.Reporting – To be recorded in a blue lab book
9.1
Sample notes (i.e. Date, UCID, AID, Sample type (i.e. Abo. Digest) and Sample
Number) should all be recorded before starting procedure
9.2
Concerns regarding wash procedure
- If no concerns; Sample washed as per SOP: No Concerns
9.3
Number of larva accounted for (and reserved if different)
10.Regulatory / Standards
N/A
11.Trouble Shooting
N/A
Appendix ll
Variable
|
Obs
Mean
Std. Dev.
Min
Max
-------------+------------------------------------------------------------Herd
|
41
2
0
2
2
Age
|
41
5.947561
2.517984
2.85
11.85
-------------+------------------------------------------------------------Pregnancy status
|
41
.6829268
.471117
0
1
Lactation status
|
41
.195122
.4012177
0
1
Body Weight
|
41
56.47561
7.676222
38
69.5
-------------+------------------------------------------------------------Backfat depth
|
41
6.02439
7.164104
0
25
-------------+------------------------------------------------------------Riney index
|
41
54.13171
25.83972
18
136.5
-------------+------------------------------------------------------------Marrow fat%
|
41
85.7122
3.708247
74.1
90.8
-------------+------------------------------------------------------------# Larvae
|
18
637.2222
410.2339
265
2000
# Males
|
18
33.33333
23.94847
5
85
# Females
|
18
50.27778
35.82934
5
110
# Bits
|
15
21.66667
16.10974
5
50
-------------+-------------------------------------------------------------