Immunology Letters 162 (2014) 69–76
Contents lists available at ScienceDirect
Immunology Letters
journal homepage: www.elsevier.com/locate/immlet
Review
Rumen transfaunation
E.J. DePeters a,∗ , L.W. George b
a
b
Department of Animal Science, University of California, Davis, CA, USA
School of Veterinary Medicine, University of California, Davis, CA, USA
a r t i c l e
i n f o
Article history:
Available online 26 September 2014
Keywords:
Rumen transfaunation
Rumen fluid
Simple indigestion
Microorganisms
a b s t r a c t
The aim of this invited mini-review is to summarize the rumen transfaunation literature. Rumen transfaunation using the cud from a healthy donor animal to treat a sick recipient animal was practiced long
before our understanding of rumen microorganisms. Around the mid-1900s, scientists began to explore
the benefits of rumen transfaunation and the associated microbial populations. Rumen transfaunation
has been used clinically to treat indigestion and to enhance the return of normal rumen function following surgical correction of a left-displaced abomasum. Rumen transfaunation was also used to introduce
unique rumen microorganisms into animals that were exposed to toxic compounds in plants. Rumen
liquor contains chemical constituents that likely contribute to the beneficial effects of re-establishing
a normal reticulo-rumen anaerobic fermentation. Recommendations for collecting rumen fluid, storage
and volumes transferred are discussed. Rumen transfaunation is a common practice to treat indigestion on dairy and livestock operations. The support of a healthy microbial community in the digestive
tract is also used for humans. Fecal microbiota transplantation has been used to treat digestive disorders
in humans. Rumen transfaunation, although not widely studied with respect to mode of action, is an
effective, practical, and easy method to treat simple indigestion of ruminants.
Published by Elsevier B.V.
1. Introduction
The aim of this invited mini-review is to summarize the information available related to rumen transfaunation. Even though the
method has been practice for decades [1] and is a common medical practice in food animal medicine to treat simple indigestion
of ruminants [2–8], there is a paucity of scientific information to
describe its benefits.
The ruminant can be considered to be a superorganism because
it has a symbiotic relationship of life between the cells of the animal’s body and the rumen microbes. Factors affecting the viability
of microorganisms in the reticulo-rumen as well as anywhere along
the gastro-intestinal tract of the ruminant impact the host animal.
Ruminants are mammals (Class – Mammalia) in the Order Arteriodactyla (even toed, hooved mammals) Suborder Ruminantia.
Ruminant, from the Latin word ruminare, means to chew over gain
hence the designation of cud chewing. Ruminants have a stomach
with four compartments or chambers – reticulum, rumen, omasum, and abomasum (Fig. 1). The reticulum, rumen, and omasum
∗ Corresponding author at: Department of Animal Science, University of California,
1 Shields Avenue, Davis, CA, USA. Tel.: +1 530 752 1263; fax: +1 530 752 0175.
E-mail addresses: ejdepeters@ucdavis.edu (E.J. DePeters),
lwgeorge@ucdavis.edu (L.W. George).
http://dx.doi.org/10.1016/j.imlet.2014.05.009
0165-2478/Published by Elsevier B.V.
are lined with non-glandular mucous membranes while the abomasum, the gastric compartment, is lined with glandular mucosa.
The abomasum is similar in function to the human stomach. The
largest compartment is the rumen, which along with the reticulum, serve as sites of anaerobic fermentation. There are coronary
grooves in the rumen creating sacs. There is a cranial groove that
separates the reticulum and rumen, and in cattle, sheep, and goats
the two compartments are easily distinguished with the reticulum having a honey combed appearance. These compartments
are lined with finger like projections called papillae that absorb
nutrients (e.g. volatile fatty acids produced by the rumen microbiota). These finger-like projects are nature’s way of increasing the
absorptive surface area of the reticulum and rumen. Often ruminant
nutritionists refer to these compartments as the reticulo-rumen
because together they function in the rumen cycle (coordinated
contractions) to support the acts of eructation and rumination. The
contractions of the rumen cycle inoculate new food with microorganisms, distribute the end products of digestion for absorption
by the mucosa papillae, and pass digesta to the omasum. Eructation is the process by which ruminants release gases from the
reticulo-rumen that are produced during anaerobic fermentation.
Eructation is a quiet process that involves eructated gas passing up
the esophagus and into the trachea and the lungs to be respired.
Rumination involves bringing a bolus of digesta up the esophagus
(regurgitate) into the mouth where the bolus of digesta (cud) is
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Fig. 1. Ruminant stomach. http://biobook.nerinxhs.org/bb/systems/digestion/
1000px-Abomasum-ia-omaso.svg.png.
chewed. The cud is eventually re-swallowed and the process continues with another bolus regurgitated for cud chewing (crushing
and grinding of particles by the molars). Cud chewing increases
surface area of the feed particles, in particular fibrous material, to
enhance microbial digestion. The act of cud chewing also stimulates salvia production, and the buffers present in the saliva help to
maintain rumen pH when the bolus is re-swallowed. Digesta leaves
the reticulum via the reticulo-omasal orifice. The omasum, with its
many leaves or laminae, controls flow of digesta to the abomasum.
The abomasum is the gastric, glandular compartment similar to
the stomach of nonruminants (human, pig, mouse) with secretion
of acid (HCl) and pepsinogen and a pyloric sphincter that regulates
flow of digesta from the abomasum to the duodenum.
Transfaunation in its current use includes a broad spectrum of
microorganisms including bacteria, protozoa, fungi, and archaea
that are transferred from rumen of a donor to the rumen of a recipient. In his seminal book on rumen microbiology, Hungate [9] stated
that transfaunation of protozoa occurred when protozoa that were
left on food by one animal were consumed by another animal.
Young animals were faunated by their mothers when she licked
them. It is not clear based on our reading of the literature where
the term ‘transfaunation’ originated or how it was derived.
One dictionary definition of transfaunation is “transfer of symbiotic fauna (usually mutualistic protozoa) from one host to another”.
Originally protozoa were defined as unicellular protists. Some ciliated protozoa have the ability to move similar to animals. Protozoa
are also eukaryotic while bacteria are prokaryotic. Defaunation of
the rumen referred to elimination of protozoa [9]. One dictionary
definition of defaunate is “elimination of microscopic fauna, especially protozoa, in the rumen and cecum, with depressing effects on
digestion”. Although the meaning of transfaunation is not clearly
defined in the literature, in this review we will use the term transfaunation broadly. The discussion of rumen transfaunation will
include both microflora and to a limited extent chemical constituents in the rumen contents.
2. Background
Brag and Hansen [1] stated rumen transfaunation was used
long before research demonstrated the importance of the rumen
microorganisms to animal metabolism. These authors reported that
the earliest printed reference about transfaunation in Sweden that
they found was from 1776 (Hjortberg) that stated “It is common
practice, even in the country side, to take the fodder out of the mouth
of a sheep or a goat to give it to an animal which does not ruminate.”
Subsequent research related to the discovery of the importance of
rumen microbial population occurred much later in the 1900s.
Research that was conducted at The Ohio Agricultural Experiment Station determined that the cud inoculated rumens of
preweaned calves contained bacteria and protozoa as early as 3
weeks of age [10]. Rumens of non-inoculated control calves that
were fed milk and alfalfa hay had only bacteria. A similar response
in calves was observed when the cud material was obtained from
cows grazing pasture [11]. Cud inoculated calves also digested a
higher proportion of cellulose and dry matter compared with noninoculated controls [12]. This advantage disappeared later once
calves were fed an all forage diet. Presumably this loss of digestion advantage with inoculation was associated with normal rumen
development of microflora at weaning. Rumen inoculation was
subsequently used as a treatment method to impact calf health. In
a field study with a herd experiencing bloody diarrhea and death of
preweaned calves, rumen transfaunation improved calf health and
survival [13].
Rumen fluid [14] from an alfalfa hay fed steer was transferred
into protozoa free sheep that were fed either alfalfa (n = 3) or a
high concentrate diet (n = 3). All 24 species of protozoa were established in the rumen of sheep fed alfalfa but only 9 species were
established in rumen of sheep fed concentrate. Rumen protozoa
play an important role in transfaunation. Rumen protozoa are predominately ciliates of two types: Entodiniomorphid protozoa and
Holotrichs [15]. Garry [16] noted that rumen protozoa were sensitive to pH, which supports the lower number of protozoa when
sheep were fed a concentrate diet [14]. Most rumen ciliates utilize
starch and their numbers increased [15]. Feeding starch that caused
a decrease in rumen pH reduced or eliminated rumen protozoa with
the larger Holotrichs more sensitive to low pH. However, it is not
simply the starch content of the diet that impacts rumen pH and
protozoa numbers, but also the type of starch and its rumen availability, the fiber content of the diet, and the physical form of the
fiber source, as well as other factors [15,17]. These dietary factors
should be considered with respect to the rumen environment of
both the donor and recipient animals when rumen transfaunation
is performed.
3. Rumen transfaunation for digestive disorders
3.1. Simple indigestion
A clinical sign of simple indigestion in dairy cattle is anorexia
(reduction in appetite) [18] with ruminal hypomotility to atony
(stasis) [6]. Sudden changes in dietary ingredients may initiate
anorexia in ruminants [4] that are reflected in changes in rumen
pH [6]. For example, changes in dietary ingredients that contribute
to rapid lactic acid production impact rumen pH and populations
of rumen microorganisms, in particular a decrease in rumen protozoa with increase acidity. Steen [19] provided diagnostic criteria
for indigestion that included (1) ketotest (test for ketones) of 0 or
1 and (2) one of the following rumen fluid parameters including
(a) a methylene blue reduction time > 3 min, (b) few large or small
protozoa, or (c) reduced protozoal activity.
Even though transfaunation of rumen fluid from a healthy door
animal to an animal with simple indigestion is a common recommended practice for dairy cattle and other ruminants, there is little
information on the practice in the scientific literature. Jasmin et al.
[20] reported the beneficial effects of rumen transfaunation for
sheep used in biomedical research that developed simple indigestion. In their biomedical research sheep were fed pelleted diets,
which contributed to the development of subclinical rumen acidosis. Exacerbating the effects of the small particle size as well as grain
content associated with the pelleted diet, there were also stresses
E.J. DePeters, L.W. George / Immunology Letters 162 (2014) 69–76
associated with shipping and indoor housing, increased handling,
fasting for experimental procedures, and perioperative opioids for
pain management – all factors that were implicated by the authors
with contributing to rumen atony and the manifestation of subclinical acidosis. To mimic the effect of indigestion, sheep were
fed a completely pelleted diet. Postoperatively two sheep on the
pelleted diet displayed rumen atony, feed intake was reduced by
greater than 50%, and protozoal motility was reduced by 25–50%.
Rumen transfaunation of one sheep once with 750 mL of fresh
rumen contents returned the recipient animal to normal health
although the other recipient animal required a second transfaunation with 750 mL of rumen fluid 24 h after the first treatment before
returning to normal. Unfortunately, these authors did not measure
rumen pH and volatile fatty acids post-transfaunation of the two
sheep that recovered from simple indigestion. Regardless, rumen
transfaunation corrected simple indigestion in sheep. It is common
knowledge in the popular literature that rumen transfaunation for
simple indigestion is effective treatment [21].
Simple indigestion can be an issue during early lactation of the
transition period when dietary changes require dairy cows to adjust
to diets high in nonstructural carbohydrates. During the transition
period around calving cows experience not only diet changes, but
also physiological changes associated with parturition (process of
giving birth). Cows are also moving from different pens so social
structure is changing and this can be a stressor. Tankersley et al.
[22] used rumen transfaunation in the transition period of dairy
cows in a field study with 210 Holstein dairy cows. Four ruminally fistulated, lactating dairy cows receiving a total mixed-ration
of concentrate and forage were used as donor animals. These four
ruminally fistulated cows were managed similar to all other lactating cows and were provided no special considerations. Rumen fluid
transfaunation was one of four treatments applied where 11.4 L of
rumen fluid were transferred by oral stomach-tube into the rumen
of fresh cows (donor cows) approximately 24-h post calving. Treatments were (1) control – no oral supplement, (2) warm water via
stomach tube, (3) commercial product via stomach tube, and (4)
rumen fluid via stomach tube. The hypothesis was that rumen fluid
transfaunation would improve health of cows after calving. Compared with control, all oral treatments did not affect serum analytes,
milk yield, and animal health in a well-managed herd. However,
a similar study should be done in a herd experiencing health and
performance issues to evaluate the impact of rumen transfaunation.
3.2. Left sided abomasal displacement
Rumen transfaunation was used as an adjunctive treatment
following surgery. Rager et al. [23] transfaunated cows following
surgical repair of left-displaced abomasum (LDA), a condition that
is a 180-degree torsion of the abomasum occurring without volvulus (no restriction of digesta passage out of the abomasum). Rumen
fluid (10 L) obtained from two non-lactating, ruminally fistulated
donor cows fed a forage diet (e.g. diet composed of predominately
hay) was transferred by stomach tube (oral) within 20 min of rumen
fluid collection. Control cows received 10 L of lukewarm tap water
also by stomach tube. Treatments occurred immediately following
surgery and again on day 1 after surgery. Beginning on day 2 following surgery and for the next three days, rumen transfaunated
cows had higher dry matter intake and milk yield compared with
control cows. Rumen fluid pH and total concentration of volatile
fatty acids did not differ for day post-surgery or between rumen
transfaunated and control treatments. Serum concentrations of ␤hydroxybutyrate on days 3 and 5 post-surgery were significantly
lower in transfaunated cows than control cows. The authors concluded “administration of rumen fluid to cows convalescing after
surgical correction of LDA had beneficial effects”.
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4. Recommended method for rumen transfaunation
Recommendations in the literature vary for rumen transfaunation, but a brief summary follows.
Rumen transfaunate can be obtained during abattoir slaughter or at post mortem. However, using a non-screened donor
may introduce infectious agents including Mycobacterium avium
var paratb, Salmonella, Cryptospordia, or Escherichia coli (0157:H7).
Other methods of transfaunate collection include stomach tube
(oral or nasal passage of the tube), cud transfaunation, or direct
removal from a donor with a surgically implanted rumen fistula
[3–5,7]. Intubation and cud transfers provide limited volume, but
may be preferable to having a known disease free source over
a random abattoir collected sample. The efficacy of using a cud
to transfaunate an adult animal is unknown, and may not provide sufficient microorganisms and substrates for full therapeutic
benefit [4]. Mould et al. [24] cautioned that the microbial quality of rumen fluid collected prior to slaughter could be impacted
by the fact that animals are often restricted in water and feed
intake. The clinical implications of fasting related changes in rumen
fluid are unknown. Radostits et al. [18] suggested that 20–30 L of
water should be pumped into the rumen via stomach tube first followed by allowing the rumen fluid to flow out of the animal by
siphon through the tube. Typically rumen fluid is transferred to
the sick animal via a stomach tube, but an oral drench can be used
[18].
Collection of large volumes of rumen fluid is most easily accomplished using a rumen fistulated animal. Removal of rumen fluid is
less stressful when the donor is fistulated than by stomach tubing
a non-fistulated donor. A rumen fistulated donor can be restrained
with a halter while use of a stomach tube usually requires the nonfistulated donor to be restrained in a chute. Typically fistulated
animals are housed and managed within the herd so there is limited
risk of introducing unknown diseases and the rumen fluid reflects
the diets fed.
Volume of rumen fluid transferred ranges from 1 L for calves and
16 L for adult cattle [2–5,16]. For adult cattle 8–16 L was considered
“ideal” [2,4,16]. Volume of transfaunate in clinical trials with adult
dairy cattle ranged between 10 and 11.4 L per oral dose [22,23].
Smaller volumes ranging between 1 and 4 L could be used for goats
and sheep (authors’ recommendation) while approximately 1 L was
recommended for sheep [6].
Rumen fluid should be transferred as soon as possible postcollection [5]. Some authors suggested that the rumen fluid should
be transferred to the recipient animal within 30 min of collection
[7]. But others suggested that rumen fluid can be stored for up to
9 h at room temperature and for 24 h at refrigeration temperature
[3,5]. Radostits et al. [18] reported that rumen fluid can be maintained at room temperature for several days. Lyophilized rumen
fluid was used successfully in culture media [25], but freeze drying
would not likely be a practical on-farm approach. Mould et al. [24]
reported that previous work demonstrated that failure to maintain
an anaerobic environment resulted in a decrease in ciliate protozoa
numbers and a decrease in cellulolytic and amylolytic activity of the
rumen fluid. However, if a container is sealed to reduce exposure
to air, pressure could increase CO2 in the fluid and decrease pH.
Hervás et al. [26] evaluated storage conditions for rumen fluid collected from sheep using in vitro methods. Storing rumen fluid under
CO2 at 0 ◦ C for 3–6 h did not affect fermentation characteristics. In
contrast, storing at 0 ◦ C for 24 h decreased fermentation characteristics. However, it should be noted that these researchers collected
rumen fluid from sheep after an overnight fast so the restriction of
feed and water could have impacted the quality of the rumen fluid
used. In general, the sooner the collected rumen fluid is transferred
from the donor to recipient the better. Best practice would be to
transfer the rumen fluid as soon as possible.
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The pH of the rumen fluid should be 5.5 or greater [7] and preferably 6.0 or greater. Rumen protozoa are decreased at low pH [15].
Typically rumen fluid pH decreased 2–4 h after feeding with high
starch diets to lactating cows [27] so avoid collecting rumen fluid
within about 4 h of feeding when the donor is receiving forage and
concentrate ingredients. With non-lactating cows fed a high forage
diet, rumen fluid collected from a fistulated cow can be done prior
to feeding or about 2–3 h after feeding since pH does not change
dramatically [28]. When using lactating cows with rumen fistulas,
rumen fluid collection prior to feeding will likely provide rumen
fluid above 6.0 pH. Lactating dairy cows eat throughout the day,
but typically the largest consumption occurs after milking when
cows are fed so collecting rumen samples after 4 h following a
large intake of feed will minimize the chance collecting rumen fluid
below 6.0 pH. Rumen fluid can be easily checked for pH using either
a portable pH meter or pH paper. On average the pH of rumen fluid
will be higher if the animal is consuming an all forage diet compared
with a diet of forage and concentrate. Forage type and particle size
will also have an impact when comparing hay versus silage fed to
the donor animal. Mould et al. [24] discussed timing of rumen fluid
collection relative to use of the fluid for in vitro inoculum, and many
of these considerations apply to rumen transfaunation.
Filter or strain large particulate matter from the rumen fluid
using cheese cloth [3] or a large screen (authors’ recommendation)
to remove large particles that can plug the stomach tube. The filtered material should contain small particles and their associated
attached bacteria, protozoa, and fungi.
Color of the rumen fluid will vary with diet. Garry [16] described
the rumen fluid as olive to brownish green when cattle are eating
hay and yellowish brown when cattle are eating silage or grain.
Avoid using rumen fluid that is frothy or foamy (authors’ recommendation).
Microscopic examination of protozoa can be performed if
desired. Protozoa are large enough to see with a transition microscope and the motility of the ciliated protozoa is readily apparent.
Methylene blue reduction time can be measured [4] by adding
0.5 mL of 0.03% methylene blue to 10 mL of ruminal fluid in a test
tube followed by mixing. The rumen environment is highly reduced
(high [H+ ] concentration). The blue color of the methylene blue
should disappear within 2–6 min [4] and if the time require is
greater than 10–15 min, it might be best to discard this rumen fluid
(Fig. 3). A robust microbial population will reduce methylene blue
to a colorless form.
For cattle experiencing simple indigestion, in addition to rumen
transfaunation, offer grass hay or straw because these are preferred
over alfalfa hay or concentrate [4]. The experiences of the authors
of this review are that cattle prefer oat hay over alfalfa hay when
experiencing simple indigestion.
Rumen fluid can be easily collected from a ruminally fistulated
donor animal by creating a siphon (Fig. 2). The collected rumen fluid
is best manually pumped into the recipient using a large polyethylene or metal stomach tube and a marine water pump. There are
commercially available tubes and pumps available with a common
pump system called “The Magrath Cattle Pump System”.
5. Rumen transfaunation in a research setting
Williams and Withers [29] studied the re-introduction of ciliated protozoa following rumen transfaunation of defaunated
sheep. Within 11 days of rumen transfaunation, ciliate protozoa
recolonized the rumens of all sheep. Even though protozoa are
reported to decrease bacterial populations due to competition for
nutrients and predation, bacterial as well as fungal populations
were not affected by the recolonization of the rumens by protozoa.
Imai et al. [30] transferred rumen fluid from Japanese Sika deer that
Fig. 2. Collection of rumen fluid from a rumen-fistulated donor and subsequent
transfer of strained rumen-fluid to a recipient animal.
E.J. DePeters, L.W. George / Immunology Letters 162 (2014) 69–76
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It required many days, 61 days for one recipient, for the bacterial
community composition to return to preexchange.
6. Application of rumen transfaunation for plant toxicants
Fig. 3. Methylene blue test.
contained nine species of ciliate protozoa into unfaunated shorthorn calves. All species of ciliates were present in calves by 14-days
post transfaunation.
Weimer et al. [31] ‘exchanged’ greater than 95% of the ruminal contents of the recipient animal with host contents to study
bacterial communities. Within about 24 h the pH and total VFA
concentrations in the recipient returned to preexchanged values.
An interesting application of rumen transfaunation is the
story related to the degradation of mimosine (␣-amino-␤-(N-[3hydroxy-4-pyridone]) propionic acid). Mimosine is a toxic amino
acid found in plants of the genera Leucaena and Mimosa [32,33].
Mimosine inhibits protein synthesis and when consumed longterm by animals, it resulted in reduced growth and loss of hair
with presumed antimitotic activity [33]. Mimosine was degraded
in the rumen to 3-hydroxy-4-pyridone [33], and in mice and rats
this metabolite of mimosine was a goitrogen [34].
Mimosine is present in leguminous plants that grow in Australia
and Hawaii [32]. Cattle and goats grazing in Australia had the
rumen microflora to degrade mimosine but not its toxic metabolite, 3,4-dihydroxy pyridine. In contrast, leucaena was not toxic to
ruminants in Hawaii and Indonesia because the rumen microflora
could degrade both mimosine and 3,4-dihydroxy pyridine [35].
Jones and Megarrity [32] isolated and cultured microorganisms
capable of degrading 3,4-dihydroxy pyridine from the rumen of a
goat in Hawaii. A bacterial culture was developed and then taken to
Australia where it was infused into one goat via a rumen fistula and
one steer via a stomach tube. The diet of both animals contained
leucaena and both animals prior to the infusion had 3,4-dihydroxy
pyridine in their urine. Following the infusion of the bacterial culture, both animals had essentially no 3,4-dihydroxy pyridine in
their urine and in vitro studies with rumen fluid obtained from the
goat demonstrated that 3,4-dihydroxy pyridine was degraded. In a
subsequent study by Quirk et al. [35] cattle were either dosed with
a bacterial culture capable of degrading 3,4-dihydroxy pyridine or
left untreated while grazing pastures containing leucaena. Dosed
steers had greater live weight gain by week 19 of the study. However, after 19 weeks, the introduced bacteria were present in the
untreated cattle. After week 19 the untreated cattle had low levels of 3,4-dihydroxy pyridine in their urine and their rumen fluid
degraded 3,4-dihydroxy pyridine in vitro. Even though the pastures
were designed to reduce the chance of transferring the introduced
bacteria to the untreated group, transfaunation occurred. These scientists attributed the transfaunation of the untreated animals to the
possibility of airborne spores from the feces of treated animals.
More recently rumen fluid from goats with rumen bacteria capable of degrading sodium monofluoroacetate, a toxic compound in
Amorimia spp., was used to transfaunate susceptible animals as a
method to reduce animal poisoning [36]. Sodium monofluoroacetate was a possible link to sudden death in Brazilian cattle that
consumed toxic plants. An interesting aspect of monofluoroacetate
is that it inhibited methanogenesis in an anaerobic sample of rumen
fluid [37].
The diversity of microbes and their ability to detoxify plant toxins will continue to be of interest in the future. Tannins are a group
of polyphenolic compounds found in plants. Tannins are antimicrobial to some species of microorganisms and were shown to reduce
methane production in sheep and goats [38]. Tannin consumption
can also have harmful effects on animal health [39]. Tannins bind
proteins and efforts are underway to use this binding property to
reduce protein degradation in the rumen to enhance nitrogen utilization by ruminants [40] and to reduce bloat of cattle grazing
alfalfa [38]. However, if tannins are to be effective it might require
that they be degraded in the rumen. Rumen bacteria were isolated
from several wild East African ruminants that were tolerant in culture to tannin [41]. Rumen cultures of selected domestic and wild
ruminants in Ethiopia partially degraded tannins in in vitro cultures [42]. These researchers also isolated fecal microorganisms
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from dikdik that completely degraded hydrolyzable tannin. Fecal
microorganisms from a number of wild ruminants had the ability
to completely degrade tannins [43]. Transfaunation of ruminants
grazing plants containing tannins with tannin degrading microorganisms might be a potential approach to mitigate the impact of
tannins on feed intake.
Even though there are situations of establishment of bacteria in
the rumen to deal with specific plant compounds, Weimer et al.
[31] reported that attempts to introduce specific bacterial strains
into the rumen have been, for the most part, unsuccessful due to
host specificity.
7. Rumen fistula techniques
There are numerous publications on the surgical procedures
involved in rumen fistulation and we cite only a few [5,44–49].
Detailed instructions with photographs are published as an internet
document.
http://www.bardiamond.com/Library.php.
http://www.bardiamond.com/Surgery Animal Care.html.
http://www.bardiamond.com/uploads/Rumen Fistula SurgeryCattle Bar DiamondTM.pdf.
The procedure is relatively simple and authors of this paper
have done numerous rumen fistulations in cattle, sheep, and goats.
Basically a circular incision is made into the skin and external
abdominal oblique muscle. The circular part of the skin and muscle
are removed. This is followed by a grid incision into the internal
oblique and transversus abdominis muscle in cattle. The peritoneum is incised sharply. The rumen wall is exteriorized and is
sutured to the muscular walls and peritoneum using overlapping
mattress sutures. The rumen wall is then sutured to the skin edge,
and the wound is covered with sterile gauze for 72 h. After that time
the circular part of the rumen wall is removed and the cannula is
inserted. However, it is also possible to immediately remove the
circular part of the rumen wall once the rumen wall is sutured to
the skin edge and then insert the cannula.
One major concern during surgery is keeping the fistula opening
size so the cannula fits snugly. If the fistula opening is too large,
the cannula can fall out of the rumen or in some cases it can fall
into the rumen. If it falls into the rumen, the cannula can often
be found in the reticulum. If the cannula falls out a less flexible
inner washer can be used. Tankersley et al. [22] used four ruminally
fistulated, lactating dairy cows on a 1500 cow (milk herd size) dairy
with no problems. The rumen fisulated cows were managed similar
to any other cow in the herd with no special considerations. The
current cannulas are made of soft material and designed to come
out if a cow should catch the cannula on an obstruction, for example
a cable. The cannula is easily re-inserted into the fistula. At the
U.C. Davis dairy facility, the rumen cannulas are routinely replaced
about every 18 months in the fistulated cows since they begin to
break down with age and exposure to sunlight. The surgeries [22]
were performed by the herd veterinarian in less than 60 min per
animal.
Point of interest: Loosli [50] stated that “rumen fistulas were first
mentioned by Fluorens in 1833”. However, the first animal to be
reported in the literature with a fistula into the stomach compartment might have been a human by the name of Alexis St. Martin
[45]. The story various depending on the source, but one version
is that in June of 1822, an army surgeon by the name of William
Beaumont was called to treat a French Canadian voyager, Alexis St.
Martin, who was shot accidently with a shotgun. St. Martin survived
but the skin and stomach wall had healed creating a permanent
opening, fistula, into his stomach compartment. Over the next 8
years, Beaumont conducted studies on gastric physiology. Many of
the aspects of gastric physiology that we know today come from
the work of Beaumont. For example, the stomach was observed to
secrete acid in response to the presence of food in the stomach and
not produced constantly as was the thought at that time. Beaumont
also noted that mucous was secreted and secreted separately from
acid. Beaumont conducted over 100 different studies on St. Martin, who lived into his 80s (depending on source maybe 86) and he
fathered numerous children (depending on source maybe 17). The
stomach fistula did not impact St. Martin’s life span as far as we
know, and the same happens for rumen fistulated dairy cattle that
typically live beyond 10 years of age at U.C. Davis.
8. Intrinsic aspects of transfaunate
Rumen fluid contains many chemical constituents that likely
contribute to the beneficial effects of transfaunation. Such factors
could include volatile fatty acids, bicarbonate buffers, proteins, and
intact microbes although there are many unidentified constituents
in rumen fluid. The effects of mechanical stimulation by bulk activity of the transfaunate may also be beneficial.
Moderate mechanical distension in the area of the reticulum
and reticulo-rumen fold can excite tension receptors [51] located
in the walls of the rumen and reticulum [52]. Leek [51] stated that
there could also be “unknown chemical stimulation arising from
fermentation” that caused muscular contractions. Transfaunation
with 8–16 L of rumen fluid in cattle could induce mechanical stimulation of tension receptors to stimulate rumination, salvation, and
normal rumen motility. The effects of osmolal changes in recipients
following transfaunation are unknown.
Bryant and Doetsch [53] demonstrated that Bacteroides succinogenogenes, cellulolytic bacteria, required factors in rumen fluid
for growth. These factors were branched-chain and straight-chain
saturated acids. Subsequent research demonstrated that ammonia
was essential for growth even in the presence of amino acids [54].
These researchers also found that biotin and selected minerals were
necessary for growth of B. succinogenogenes. Streptococcus bovis ferments starch, is acid tolerant, and can grow rapidly. Under “normal
conditions” S. bovis produced acetate, formate, and ethanol [55]. In
response to excess readily rumen digestible carbohydrates, S. bovis
yielded lactate, which contributes to subclinical acidosis associated
with diet changes and indigestion. S. bovis required biotin, thiamine
was stimulatory, and some strains were reported to require the B
vitamins, pantothenic acid and nicotinic acid [9]. Ammonia could
serve as the nitrogen source. There is a cross feeding of nutrients
among the microorganisms that live in the rumen [56,57]. Miura
et al. [57] described it as nutritional interdependence where, for
example, the branch-chain fatty acid produced by one bacterial
group was used by branch-chain fatty acid dependent groups for
growth. There was reported to be considerable cross feeding of
cellulose degradation products in the rumen [58]. Likewise, the B
vitamins produced by some organisms are required for growth by
other organisms. These are just a few examples of how complex
the rumen environment is and how interlinked the microorganisms can be. The nutrients in rumen fluid transfaunate will likely
benefit the growth of bacteria, protozoa, methanogens, and fungi
to restore normal microflora populations.
Protozoa in the rumen transfaunate likely play an important role
in re-establishing the rumen microbial population. Protozoa concentration increased as the proportion of starch or sugar in the diet
increased [59]. Protozoa were reported [59] to stabilize the rumen
pH in response to the consumption of readily available starch and to
decrease the redox potential. Consequently these actions create a
rumen environment conducive to cellulolytic bacteria growth. Protozoa also degrade protein and contribute peptides that stimulate
the growth of other microorganisms.
E.J. DePeters, L.W. George / Immunology Letters 162 (2014) 69–76
The intrinsic factors associated with rumen fluid are not
well described or understood, but there are obvious benefits to
the animal. Technology is now available to study ruminal fluid
metabolome [60] and improve our understanding.
9. Fecal microbiota transplantation
Transfaunating the rumen of sick animals using the microorganisms obtained from a healthy donor is common practice. Microbial
transfer is also used in human medicine. The transfer of beneficial fecal microorganism (fecal microbiota transplantation; FMT)
was reported to be practiced for more than 40–50 years [61,62].
The concepts for ruminants and humans are similar – support a
healthy microbiology population in the digestive system and there
is a healthy host. Humans can be thought of as superorganisms
because there is a vibrant microbial population in symbiotic relation with the host. Microbial floral was re-established in patients
suffering from Pseudomembraneous enterocolitis [63]. Borody and
Khortus [62] discussed the benefit of FMT with respect to Clostridium difficile infection following the use of antibiotics that disrupted
the microbial community. Treatment of C. difficile involved antibiotics, but relapses occurred. Cure rates were high using FMT [64,65].
Following FMT the bacterial population of the recipient reflected
the bacterial population of the donor [66,67]. Bakken [68] used
what was called fecal bacteriotherapy to successfully cure C. difficile (90% success rate). The fecal liquid suspension from a donor
was transferred into a recipient “from either end of the GI tract”.
10. Summary
Rumen transfaunation is a routine, widely accepted, successful
procedure to treat simple indigestion in ruminants. The procedure
also has clinical application for post-operative treatment of cattle
with left sided abomasal displacements. Rumen fluid from a healthy
donor provides the recipient with diverse microorganisms that
can repopulate the rumen. Transplanted rumen fluid also provides
nutrients and energy to the rumen microbial population. Although
widely accepted as a treatment for simple indigestion, there is a
paucity of information in the literature to describe the mechanisms
involved in the beneficial effects of rumen transfaunation.
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