Upload
others
View
8
Download
0
Embed Size (px)
Citation preview
i
EPIDEMIOLOGY, DIAGNOSIS AND CHEMOTHERAPY OF STRANGLES IN EQUINES
By
Muhammad Ijaz 2005-VA-150
A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENT FOR THE DEGREE
Of
DOCTOR OF PHILOSOPHY
In
CLINICAL MEDICINE
Department of Clinical Medicine and Surgery FACULTY OF VETERINARY SCIENCE
UNIVERSITY OF VETERINARY AND ANIMAL SCIENCES, LAHORE
2010
ii
In the name of Lord who created, Created man from clot,
And thy Lord is the most Bounteous, Who taught by pen,
Taught man when he knew not.
iii
To
The Controller of Examinations, University of Veterinary and Animal Sciences, Lahore.
We, the Supervisory Committee, certify that the contents and form of the
thesis, submitted by Mr. Muhammad Ijaz, Regd No, 2005-VA-150, have been found
satisfactory and recommend that it be processed for the evaluation of External
Examiner(s) for the award of degree.
Chairman___________________________
Prof. Dr. Muhammad Sawar Khan
Member_____________________________
Prof. Dr. Muhammad Arif Khan
Member ______________________________
Prof. Dr. Azhar Maqbool
iv
WISDOM IS THE PART AND PARCEL
OF MY RELIGION,
KNOWLEDGE MY WEAPON,
PATIENCE MY DRESS,
FAITH MY DIET, AND SINCERITY
MY COMPANION
HADIS – E – NABVI (PEACE BE UPON HIM)
v
I
DEDICATE THE FRUIT OF THIS HUMBLE EFFORT TO
THE HOLY PROPHET
(PEACE BE UPON HIM)
THE GREAT SOCIAL REFORMER
AND MY
MAGNIFICENTLY PRECIOUS
BELOVED PARENTS, MY SUPERVISER, MY BROTHERS & MY FRIENDS
WHO ALWAYS APPRECIATE AND PRAY FOR ME TO ACHIEVE HIGHER GOALS OF LIFE.
vi
AACCKKNNOOWWLLEEDDGGEEMMEENNTTSS
I am thankful to most gracious and ALMIGHTY ALLAH (AZAWAJAL)
who gave me the health and opportunity to complete this work, I bow before my
compassionate endowments, Peace be upon HOLY PROPHET MUHAMMAD
(PEACE BE UPON HIM) who is ever an ember of guidance and knowledge for
humanity.
I feel great honor to place on the record my sincere thanks to my
supervisor Prof. Dr. Muhammad Sarwar Khan, Department of Clinical Medicine
and Surgery, University of Veterinary and Animal Sciences, Lahore. He
supervised my research lightheartedly and proficiently made the dispatch of
intimidating work load possible by persistent guidance and scholarly criticism
communicated to me during the course of this study and execution of this
manuscript.
The co-operation extended by the members of my supervisory committee,
Prof. Dr. Muhammad Arif Khan, Chairman, Department of Clinical Medicine
and Surgery, University of Veterinary and Animal Sciences Lahore and Prof. Dr.
Azhar Maqbool, Department of Parasitology, University of Veterinary and
Animal Sciences, Lahore is very sincerely appreciated for their skillful
suggestions during the whole span of this investigation.
I wish to express my gratitude to Prof. Dr. John F. Timoney, for his
guidance, advice and support during this work. I am very thankful to him for
providing me the research facilities which helped in timely completion of this
work. I would like to thank the valuable contribution of Dr. Sridhar Vilineni. I
also wish to thank my lab members, Dr. Sergey Artiushin and Mike Fettinger for
their valuable input.
I have great sense of obligation to reverend Dr. Muhammad Muddassir
Ali, Dr. Muhammad Avais and Dr Muhammad Hassan Saleem, Department of
vii
Clinical Medicine and Surgery, University of Veterinary and Animal Sciences
Lahore, for their keen interest, propitious guidance, enlightened views and
valuable suggestions for the successful accomplishment of present study. I am also
thankful to the Laboratory staff Department of Clinical Medicine and Surgery.
I am whole heartedly thankful to Dr. Muhammad Arshad Shad (Lt. Col)
being my elder brother, his keen interest and indefatigable help with anything,
anytime, any where and computing exhaustive statistical evaluation from raw
data, made it all possible for me to undertake and complete this project
successfully.
I am extremely grateful to Higher Education Commission of Pakistan for
providing me financial support for this study.
I affectionally like to revive and appreciate the sincerity, help and
encouragement of my friends Dr. Zafar ullah Khan, Dr. Arslan Farooq, Dr. Agha
Shahzad, Dr. Zeshan M. Iqbal, Dr. Sohail Akbar, Dr. Muhammad Farooq, Dr.
Asim Munawar, Dr. Abdul Rehman, Dr Umair Iftikhar, Dr Manuchahar Ali, Dr.
Tanveer Hussain, Dr. Mir Ahmad, Dr Abdul Qadus, Dr. Khalid Mehmood, Dr.
Ahmad Jawad Sabir, Samuel Shahzad, Dr. Salman Khalid and all of other friends.
Last but not least, I must acknowledge my indebtedness to my loving
parents, and brothers for the motivation to take up this program of studies,
financial support, and their hands in prayers for my success, and great patience
and goodwill throughout the fairly long period of training at this institution.
Finally, I hope that inadvertent errors will be forgiven by the readers.
MUHAMMAD IJAZ
viii
TABLE OF CONTENTS
Dedication………………………………………………………........v
Acknowledgements ……….……………………………………..…..vi
List of Tables .……..………………………………………………...ix
List of Figures……………………………………………………….xiv
SR. NO. CHAPTERS PAGE NO.
1 INTRODUCTION 1
2 REVIEW OF LITERATURE 7
3 MATERIALS AND METHODS 32
4 RESULTS 46
5 DISCUSSION 127
6 SUMMARY 145
LITERATURE CITED 151
ix
LIST OF TABLES
Table No. Title Page No.
3.1 Details of all the primers used in the study 37
3.2 Geographic origin, year of isolation and strain characteristics of S.equi
40
4.1 Prevalence of Strangles in nasal discharge and pus samples of sub-mandibular lymph nodes of horses on the basis of culture.
48
4.2 Prevalence of Strangles in nasal discharge of horses on the basis of culture.
50
4.3 Prevalence of Strangles in pus samples of sub-mandibular lymph node of horses on the basis of culture.
51
4.4 Prevalence of Strangles in nasal discharge and pus samples of sub-mandibular lymph nodes of horses on the basis of PCR.
52
4.5 Prevalence of Strangles in nasal discharge of horses on the basis of PCR.
54
4.6 Prevalence of Strangles in pus samples of sub-mandibular lymph nodes of horses on the basis of PCR.
55
4.7 Prevalence of Strangles in nasal discharge and pus samples of sub-mandibular lymph nodes of mules on the basis of culture.
56
4.8 Prevalence of Strangles in nasal discharge of mules on the basis of culture.
58
4.9 Prevalence of Strangles in pus samples of sub-mandibular lymph nodes of mules on the basis of culture.
59
4.10 Prevalence of Strangles in nasal discharge and pus samples of sub-mandibular lymph nodes of mules on the basis of PCR.
60
4.11 Prevalence of Strangles in nasal discharge of mules on the basis of PCR.
61
4.12 Prevalence of Strangles in pus samples of sub-mandibular lymph nodes of mules on the basis of PCR.
62
4.13 Mortality rate in horses under 5 years of age 64
4.14 Mortality rate in mules under 5 years of age 65
4.15 Details of all amplicons used in the present study. 67
4.16 SeM alleles in S. equi isolated over a period of 40 years in N. America, Europe and Asia
68
4.17 Identification of six new alleles by BLAST analysis against www.pubmlst.org/szooepidemicus/
70
4.18 Frequency of single nucleotide polymorphism (SNPs) in SeM, SzPSe and Se18.9 in 25 isolates of S. equi.
71
4.19 Details of SzPSe of all isolates of S. equi 72
x
Table No. Title Page No.
4.20 Comparison of culture and PCR for identification of carrier of S. equi in naturally infected horse ≤ 5 years of age.
74
4.21 Comparison of culture and PCR for identification of carrier of S. equi in naturally infected mule ≤ 5 years of age.
76
4.22 Total white blood cell count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
78
4.23 Mean segmented Neutrophilic count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
80
4.24 Total Lymphocytic count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
82
4.25 Total Monocytic count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
83
4.26 Total Eosinophil count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
84
4.27 Total Basophilic count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
86
4.28 Erythrocytes count (X 1012/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
88
4.29 Packed cell volume (%) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
89
4.30 Haemoglobin concentration (g/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
91
4.31 Total white blood cell count (x109/L) in healthy and carrier horses and mules from strangles. (Mean ± SE)
93
4.32 Mean segmented Neutrophils count (x109/L) in healthy and carrier horses and mules from strangles. (Mean ± SE)
95
4.33 Total Lymphocytic count (x109/L) in healthy and carrier horses and mules from strangles. (Mean ± SE)
96
4.34 Total Monocyte count (x109/L) in healthy and carrier horses and mules from strangles. (Mean ± SE)
98
4.35 Total Eosinophil count (x109/L) in healthy and carrier horses and mules from strangles. (Mean ± SE)
100
4.36 Total Basophil count (x109/L) in healthy and carrier horses and mules from strangles. (Mean ± SE)
102
4.37 Erythrocytes count (X 1012/L) in healthy and carrier horses and mules from strangles. (Mean ± SE) 104
4.38 Packed cell volume (%) in healthy and carrier horses and mules from strangles. (Mean ± SE)
105
4.39 Haemoglobin concentration (g/L) in healthy and carrier horses and mules from strangles. (Mean ± SE)
107
xi
Table No. Title Page No.
4.40 Total serum protein values (g/L) of healthy and diseased horses and mules suffered from strangles. (Mean ± SD)
109
4.41 Serum albumin values (g/L) of healthy and diseased horses and mules suffered from strangles. (Mean ± SD)
110
4.42 Serum globulin values (g/L) of healthy and diseased horses and mules suffered from strangles. (Mean ± SD)
112
4.43 Fibrinogen values (g/L) of healthy and diseased horses and mules suffered from strangles. (Mean ± SD)
114
4.44 In-vitro Antibiotic sensitivity against S. equi in horses 116
4.45 In-vitro Antibiotic sensitivity against S. equi in mules 117
4.46 In-vivo Antibiotic sensitivity against S. equi in horses 119
4.47 In-vivo Antibiotic sensitivity against S. equi mules 120
4.48 In-vitro efficacy of Phenol as disinfectant against S. equi by using Phenol Coefficient test
121
4.49 In-vitro efficacy of Dettol as disinfectant against S. equi by using Phenol Coefficient test
122
4.50 In-vitro efficacy of Povidone Iodine as disinfectant against S. equi by using Phenol Coefficient test
123
4.51 In-vitro efficacy of 0.6% Sulfuric acid as disinfectant against S. equi by using Phenol Coefficient test
124
4.52 In-vitro efficacy of Bleach as disinfectant against S. equi by using Phenol Coefficient test
124
4.53 Overall comparison of Different Disinfectants used against S. equi
125
xii
LIST OF FIGURES
Fig No. Title Page No.
4.1 Prevalence of Strangles in horses and mules 47
4.2 Month wise prevalence of Strangles in horses on the basis of culture. 49
4.3 Month wise prevalence of Strangles in horses on the basis of PCR. 53
4.4 Month wise prevalence of Strangles in mules on the basis of culture 57
4.5 Month wise prevalence of Strangles in mules on the basis of PCR. 60
4.6 PCR amplification of DNAs from equine isolates of S. equi with specific primers.
63
4.7 PCR amplification of DNAs from equine isolates of S. equi to analyse variation in SeM (a), SzPSe (b), Se18.7 (c) and EqbE (d).
69
4.8 Week wise comparison of Total white blood cell count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
80
4.9 Week wise comparison of Mean segmented Neutrophilic count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
82
4.10 Week wise comparison of Total Lymphocytic count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
83
4.11 Week wise comparison of Total Monocytic count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
85
4.12 Week wise comparison of Total Eosinophil count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
86
4.13 Week wise comparison of Total Basophilic count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
88
4.14 Week wise comparison of Erythrocytes count (X 1012/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
89
4.15 Week wise comparison of packed cell volume (%) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
91
4.16 Week wise comparison of Haemoglobin concentration (g/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
93
4.17 Week wise comparison of Total white blood cell count (x109/L) in healthy and carrier horses and mules. (Mean ± SE)
95
4.18 Week wise comparisons of Neutrophils count (x109/L) in healthy and carrier horses and mules. (Mean ± SE)
96
4.19 Week wise comparisons of total lymphocytic count (x109/L) in healthy 98
xiii
and carrier horses and mules. (Mean ± SE)
4.20 Week wise comparisons of total Monocytic count (x109/L) in healthy and carrier horses and mules. (Mean ± SE)
100
4.21 Week wise comparisons of total eosinophilic count (x109/L) in healthy and carrier horses and mules. (Mean ± SE)
102
4.22 Week wise comparisons of total Basophilic count (x109/L) in healthy and carrier horses and mules. (Mean ± SE)
104
4.23 Week wise comparisons of Erythrocytes count (X 1012/L) in healthy and carrier horses and mules. (Mean ± SE)
105
4.24 Week wise comparisons of packed cell volume (%) in healthy and carrier horses and mules. (Mean ± SE)
107
4.25 Week wise comparisons of Hb concentration (g/L) in healthy and carrier horses and mules. (Mean ± SE)
108
4.26 Age wise comparison of total serum protein values (g/L) of healthy and diseased horses and mules suffered from strangles. (Mean ± SD)
110
4.27 Age wise comparison of serum albumin values (g/L) of healthy and diseased horses and mules suffered from strangles. (Mean ± SD)
112
4.28 Age wise comparison of serum globulin values (g/L) of healthy and diseased horses and mules suffered from strangles. (Mean ± SD)
113
4.29 Age wise comparison of fibrinogen values (g/L) of healthy and diseased horses and mules suffered from strangles. (Mean ± SD)
116
1
Chapter-01
INTRODUCTION
According to the Agriculture Census Organization, the total equine population
in Pakistan was 4.8 million in 2006. This has risen to 5.1 million (Horses 0.4, Asses
4.5 and Mules 0.2) as per the census report for 2008-2009. (Anonymous 2008-09)
Strangles is named from the air restriction in late stages of the disease where
the horse breathes as if it is being strangled because of the restriction of the trachea
due to swollen lymph nodes. It is considered to be one of the top three most
significant and feared respiratory diseases in horses (Natarajan and Langohr, 2003). It
accounts for close to 30% of all equine infections reported worldwide, making it the
most frequently encountered single horse illness (Harrington et al., 2002).
The mechanism and route of entrance of Streptococcus equi subsp. equi (S.
equi) into the lymphoid system has not so far been properly elucidated. An in vitro
trial revealed adherence of S. equi to equine epithelial cheek cells, tongue and nasal
epithelial cells (Srivastava and Barnum, 1983b; Valentin Weigand et al., 1988).
Timoney, (1988) reported adherence of large numbers of S. equi on the soft palate
and adjacent tonsillar tissue of two experimentally infected ponies one hour post
inoculation. He further reported presence of S. equi on the soft palate, tonsils and the
retropharyngeal lymph nodes of a pony necropsy five days post infection. This
information is suggestive of sites of colonisation, subsequent to infection.
A number of researchers have investigated the link between Strangles and S.
equi. Todd, (1910) and Shultz (1888) discovered the relationship between Strangles
2
and a chain forming coccus. Bazeley and Battle (1940) found that this coccus, S. equi,
was consistently associated with strangles and Bazeley (1943) showed that
inoculation of horses with pure cultures of S. equi resulted in the reproduction of the
classic disease. Bryans et al., (1964) also confirmed the relationship in a later study.
He was also able to induce clinical disease in horses by infecting with a broth
containing S. equi.
The symptoms become apparent after an incubation period of 3 to 8 days, and
the clinical course usually lasts 3 to 4weeks (Nara et al., 1983). Marked fever (103-
106°F) develops during the acute phase and may subside until the lymph nodes
abscess, this is the time when a second wave of fever may develop. Affected horses
become anorexic, depressed, and develop bilateral, serous to mucoid nasal discharge
within 24 hours of fever. The discharge becomes mucopurulent as the disease
progresses, and a moist cough may develop in some cases. Plasma fibrinogen
concentration and leukocyte counts usually increase at this time. The submandibular
lymph nodes are involved most oftenly and become enlarged, firm, and painful. The
retropharyngeal lymph nodes may also be affected, and if they become markedly
enlarged may induce dysphagia. The abscessed lymph nodes typically rupture 7 to 10
days after the onset of clinical signs and, in uncomplicated cases; recovery is
complete 1 to 2 weeks thereafter (Reed, 2004).
The highly host adapted S. equi of Lancefield group C causes equine
strangles, a highly contagious purulent rhino-tonsillitis and lymphadenitis of the head
and neck. Isolates of S. equi constitute a clone or biovar of an ancestral S.
zooepidemicus with which it shared greater than 97% DNA homology. Its anti-
3
phagocytic SeM protein is an important protective antigen and virulence factor that
functions by binding fibrinogen thereby masking C3b binding sites on the surface of
S. equi and by directly inhibiting deposition of the opsonic forms of C3 on the
bacterial surface (Boschwitz and Timoney, 1994). Although early studies carried out
by Anzai et al. (2005) and Kelly et al. (2006) indicated that SeM was highly
conserved but variation in the N-terminal sequences from aa 37 to 143 has recently
been reported in isolates from N. America, Europe and Japan DNA codon changes
associated with non-synonymous substitutions of amino acids throughout the N-
terminus are strongly suggestive of immune selection pressure. However, allelic
variation in SeM does not significantly affect susceptibility of S. equi to opsonization
or ability to bind fibrinogen (Timoney et al., 2009). Epitopes on the variable region
reactive with convalescent mucosal IgA are more frequent than epitopes reactive with
convalescent IgG suggesting that IgA is a more significant selection pressure. There
is also evidence that N17 terminal variation affects the conformational and not the
linear epitope (Timoney et al., 2009).
SeM allelic variants of S. equi from the same animal including those with
chronic guttural pouch or cranial sinus infection have been shown to emerge over a
short period (Anzai et al., 2005; Kelly et al., 2006). Interestingly, some isolates from
the guttural pouch have large in frame deletions of the SeM N-terminus (Chanter et
al., 2000). It suggests that loss of this region has survival value in the face of local
acquired immune responses that target the N-terminus. SeM. is one of a small number
of immunoreactive 1 surface exposed and secreted proteins of S. equi that elicit
convalescent serum and mucosal antibody responses (Timoney et al., 2007). The
4
surface exposed SzPSe and secreted Se18.9 proteins are of particular interest, because
they elicit strong serum and mucosal antibody responses and bind to tonsillar
epithelial cells (Timoney et al., 2007; Tiwari et al., 2007; Fan et al., 2008). They are
therefore potentially subject to the same selection pressure as SeM during the
acquired immune response of the horse. SzPSe is a homologue of the variable Moore
and Bryans typing antigen SzP (Walker and Timoney, 1998), which has been shown
to elicit opsonic and protective antibodies. The anti-phagocytic Se18.9 is uniquely
expressed and secreted by S. equi. It also becomes associated with the bacterial
surface (Tiwari et al., 2007). Incubation of this protein with equine neutrophils causes
a significant reduction in their bactericidal activity for both S. zooepidemicus and S.
equi, an effect partially neutralized by Se18.9 specific antibody.
Once infected, the majority of animals recover and eliminate S. equi over a
period of 4–6 weeks. However, in 10% of clinically recovered cases S. equi may
continue to be shed intermittently for prolonged periods. This carrier status is
probably caused by incomplete drainage of exudate from the guttural pouches
(empyaema) and/or sinuses following rupture of abscesses formed in the
retropharyngeal lymph nodes (Newton et al., 1997).
Changes in hematological parameters, such as total and differential white cell
counts have been recorded in horses naturally and experimentally infected with S.
equi (Hamlen et al., (1994). In another study Dalgleish et al. (1993) performed blood
neutrophil count on a group of naturally infected horses. Knight et al. (1975) and
Nara et al. (1983) have also carried out hematological studies including total white
5
cell counts in horses on days 2, 5, 10, 15, and 20 following experimental infection on
horses experimentally infected with S. equi
The use of antibiotics in the treatment of strangles with early clinical signs of
infection can be of great value. By following this regime, the course of the disease is
shortened, lymph node abscesses and their complications are prevented. Degree of
environmental contamination from infected discharges and secretions is also greatly
reduced. Early identification of new cases during outbreaks, through careful
observation of exposed horses and recording of rectal temperatures twice daily, will
facilitate initiation of antimicrobial treatment early in the disease course and
maximize the effectiveness of this approach. Wilson, (1988) recommends use of
Procaine penicillin G at dose rate of 22,000 IU/kg IM twice a day for at least 10 to 14
days. He further suggests continuing it for 5 days beyond resolution of clinical signs.
The organism is also susceptible to ampicillin, ceftiofur, erythromycin, rifampin,
tetracycline, and trimethoprim/sulfonamides, although the later appears to be less
effective than beta-lactam antimicrobials in treating field cases. Inadequate doses of
antibiotics and/or an inadequate length of treatment may result in treatment failure or
recurrence. No evidence reported that early treatment with antibiotics increases the
risk for development of internal abscesses (Sonea, 1984).
Outbreaks of strangles may last for months or years within large horse
populations. It will perpetuate itself with frequent new arrivals in the horse facility
providing a continuous supply of susceptible animals (Harrington et al., 2002).
Although it is a potentially fatal disease, even then it can be contained and treated by
paying due attention.
6
Keeping in view the importance and utilization of equines in our country and
the significant losses rendered by Strangles, the present project was designed to
achieve the following objectives
To study the epidemiology of disease.
To determine whether immune selection pressure that resulted in N-terminal
sequence diversity in SeM also affected SzPSe and Se18.9 or otherwise.
PCR based diagnosis of S. equi from carrier animals.
To study the effect of Strangles on Haemogram, Leucogram and Serum
proteins.
To evaluate the efficacy of various drugs under laboratory and field
conditions.
To evaluate the efficacy of various disinfectants under laboratory conditions
7
Chapter-02
REVIEW OF LITERATURE
Van Dorssen, (1939) isolated S. equi from horses that were affected from
strangles. A total of 80 samples were collected for the study. S. equi was isolated
from the abscesses of above mentioned animals and suggested that S. equi was the
causative agent of strangles in horses and measures to be taken against this organism
to combat the disease. Bazeley and Battle, (1940) studied various types of
Strepotococci in equine infections. They isolated 457 strains of haemolytic
Streptococci from 415 cases of horses. All strains belong to group C Lancefield by
appropriate precipitin tests. On the basis of fermentation tests and colony type
Streptococci were divided into five types. The colonies of Streptococci were dom-
shaped and honey coloured and grew up to about 3 mm in diameter in 24 hours.
Tenacious strands were observed when a platinum loop was withdrawn after touching
a colony. Hignett and King, (1940) investigated the Streptococcal infection in horses
and summarized that horses are highly susceptible to Streptococci than any other
domestic animal. Streptococci group caused a wide variety of local and generalized
disease conditions in horses such as strangles, pneumonia, metritis, and sinusitis and
as commonest secondary invaders in wounds such as poll evil, fistulous withers and
quitter. Namikawa et al. (1940) used an anaerobic technique for its growth to study
the microbial characteristics of S. equi. The colony features of the bacteria presented
a “Medusa Head” formation after the 18 hours and the distinction was lost after few
hours because the bacteria growing in this form were non-encapsulated. Medusa Head
8
colonies were produced by 43 out of 44 strains of C type S. equi and 34 out of 36
strains of G type S. equi. Tuji and Sato, (1940) examined 88 cases of diseased horses
and also 6 healthy horses. From these they isolated 100 different strains of
Streptococci. They concluded that two strain of S. equi, C type strains were isolated
from a case of coryza and from strangles complication and infectious anaemia. In the
other cases, C and G type strains of S.equi were isolated from spleen and lungs
respectively. Castagnoli and Balboni, (1942) reported of S. equi in two 4 year old,
horses and two, 4 year mares in cerebrum and developed cerebral signs. From the
brain of each of the two mares S. equi was isolated. Eberbeck and Halswick, (1943)
studied the pathogenesis of strangles over a period of three years at Army Horses
under constant veterinary supervision and concluded that many of the sequelae of
strangles were of an allergic nature. Minett, (1944) studied the incidence of strangles
hardly fell below 50% in remount depot and stud farm of sub-continent during that
period 1914-1942. Mona and Sargodha are two important army remount depots in
Pakistan and reported annual incidence of strangles based on the mean annual
percentage of 1917-1940 was 76.2% at Mona and similarly the annual incidence of
strangles based on the mean annual percentage of 1927-1940 at Sargodha was 72.6%
reported. While this incidence of strangles among mules at these stations was lower,
with mean annual rates of 53.2% at Mona and 46.8% at Sargodha. When we talk
about the stud farms the yearly incidence varied from zero to 50%. Climatic factors
including rainfall, temperature, relative humidity, wind velocity and dust storms were
also considered important in occurrence of the strangles. In a more equable climate,
the attack rate was higher in the cold season than in the hot and was very much less in
9
the summer rainy season. Monsoon conditions seemed unfavorable for the spread of
strangles. Peatt, (1945) reviewed veterinary services in India and Burma in the
Second World War and reported that strangles was recorded as very virulent in two
big remount depots where the mortality was reached upto16%. Menninger, (1949)
studied the causative agent of strangles. He stated that four mares contracted typical
strangles 2-6 days and he isolated S. equi from the mandibular lymph node abscesses.
He determined that strangles in equine is due to a pathogenic bacterium, S.equi. He
also observed some predisposing factors which may either be stress of a virus
infection and or a nonspecific weakening resulting from natural causes such as fatigue
and exposure to harsh weather. Tajima and Ueda, (1953) reported his observation on
the four cases of purulent encephalomyelitis in strangles affected horses and showed
nervous signs including depression or excitement, incoordination, rigidity of the neck
during the course of strangles. Paunovic, (1957) reported an abdominal form of
strangles while having rectal examination of four horses completely with a history of
colic. Vukovic, (1961) diagnosed strangles in 114 of 1853 horses examined during
that specified period 1952-59 in Sarajevo. Out of these 114 horses only nine showed
the infection in the mesenteric lymph nodes. Mahaffey, (1962) observed that strangles
was commonest disease in large stud farms and army remounts depots where the
horses are kept together in large numbers. Strangles is highly infectious and
contagious disease of all equines and causative agent of strangles is very resistant
which may remain viable for weeks or months in dried pus. Natural infection was
acquired by direct or indirect contact between healthy, susceptible and carrier
animals, diseased animals or the products of the latter. In almost every case acute
10
inflammation of the submandibular lymph nodes commenced early during the course
of disease. The submandibular lymph nodes showed the cardinal signs of inflamation
in about 4-5 days. In the beginning they were hard but after few days they began to
fluctuate with pus inside of them. As the abscesses matured, the overlying skin
became denuded of hair and small amounts of sticky fluid oozed out over the surface.
Pharyngitis was very common along with abscessation of adjacent lymph nodes.
Roaring and some other respiratory diseases were usually seen as sequelae of
strangles. Bryans et al. (1964) mentioned the etiology of strangles. In his
experiments using 23 horses they indicated that S. equi is the only causative agent of
equine strangles. Strangles was set up by intranasal inoculation of horses with pus
from abscesses and cultures with abscesses. Wagenaar and Schaaf, (1965) discussed
the clinical picture, etiology and active immunization of strangles and concluded that
antibiotics ought not be used in cases running a normal course. In complicated cases
treatment with penicillin for 5-10 days would be useful but isolation and
identification of diseased animals is important. Wisecup et al. (1967) studied
strangles in a herd of donkeys in which the lesions consisted of mainly caseation and
calcification of the abdominal lymph nodes. S. equi was isolated from these lesions.
Culturing S. equi from mare exudates appeared to be the most reliable method for
identification of the infected animals. The clinical syndromes caused by S. equi in
donkeys showed a clear discrimination from those of strangles in horses. Roberts,
(1971) noted chorioretinitis in six mares and seven geldings after recovery from an
attack of strangles. He described the lesions grossly and histopathologicaly and
discussed their relationship with the previous infection of strangles. Woolcock,
11
(1975) studied an atypical variety of S. equi. It was shown to be deficient in capsular
material, to be very virulent for mice and possess a cell-wall protein similar to M-like
protein of classical S. equi. Antiserum prepared against classical S. equi effectively
opsonised and induced the formation of long chains by these atypical strains. It is
possible to use this variant of S. equi to overcome many of the current problems link
with the manufacture and use of strangles vaccines. Woolcock, (1975) studied the
epidemiology of equine Streptococci. Three different types of samples were collected
from equine tonsillar tissue, draining regional lymph nodes and as well as deep nasal
swabs and examined bacteriologically. Group C Streptococci, predominantly S.
zooepidemicus, was found in all types of samples. One of the most frequent sites for
isolation was the tonsil. S. equi was not found in any of the tissues sampled. Niebauer
et al. (1979) reported a case of brain metastatic strangles abscess in three year old
horse. The horse was showing signs included ataxia, swelling of the mandibular
lymph nodes and eye disorders. During post mortem examination, a tumour was
found in the subdura of the left frontal lobe and in caudate nucleus encephalitis was
established. S. equi was isolated from that tumour. Ford and Lokai, (1980) tried to
find out the method of transmission of various diseases in weanling horses. They
gathered 300 weanling horses on equal number of acres of grossly overgrazed
pasture. That pasture was previously grazed by horses, out of which some of them
had suffered from strangles. Management of the horses was so poor that when
strangles appeared; it affected all weanling horses and killed 10% of them.
Postmortem examination showed that S. equi could cause abscesses in any organ of
the body including the brain. Prescott et al. (1982) discussed a mild and weak form of
12
strangles caused by S. equi. A mild form of strangles caused by S. equi was identified
on a massive type of stud farm that was for breeding purpose. The organism varied
from most of S. equi isolates because of absence of the mucoid capsule even after 24
hours of culture, leaving a matt-type colony. Typically, the clinical signs were a
transient (24-48 hours) fever, anorexia and profuse nasal discharge. Half of the
diseased horses expressed restrained mandibular lymph node enlargement, and these
glands usually ruptured or were drained. The use of a PHAT (passive
hemagglutination antibody test) showed that subclinical infection was widespread in
horses on the farm. George et al. (1983) identified the carriers of S. equi in a naturally
infected herd. During an epidemic of strangles in a population of research horses, out
of which four mares were identified as carriers of S. equi infection. Three of the
mares were paradigm of strangles. They showed intense regional lymphadenitis with
or without rupture of abscessed lymph nodes. The 4th mare showed occurence of
serous to mucopurulent nasal discharge, but never had more than a mild degree of
lymph node enlargement. From the abscessed lymph nodes and nasopharyngeal swab
specimens S. equi was isolated from the first 3 mares from 6 to 19 weeks after rupture
of infected lymph nodes. From the nasopharynx of the 4th mare S. equi was isolated
and intermittently over the ensuing 6 months. After that, 4th mare was kept in
isolation during the 7th month, where she continued to shed S. equi for 4 more
months. An integral and meticulous physical examination during the 10th month,
including radiography of the head and thorax, did not show any pertinent
abnormalities, but a pharyngeal swab was culture-positive for S. equi. This isolate
after taking the 4th mare was used to inoculate 2 yearling colts, which developed
13
strangles and from which S. equi was reisolated. Shedding of S. equi by mare 4 was
stopped in the 11th month, and at necropsy 2 months later, S. equi was not found in
any organ or tissue. Corticosteroids administration 3 weeks prior to necropsy had
persuaded neither shedding of the S. equi nor clinical signs of strangles. This study
concluded clinical, epidemiologic, and bacteriologic documentation to support the
existence of a carrier state following natural infection with S. equi. Nara et al. (1983)
designed the experimental study of S. equi infection in horses and found out its
correlation with in vivo and in vitro immune responses. They conducted an
experiment on 14 horses, divided into 2 groups based on 18 or 24 hours skin-test
reactions to S. equi, and inoculated virulent strain of S. equi nasopharyngeally.
Animals (n = 6, group I) with confirmation of previous exposure to S. equi, with one
exception, developed very few or no signs of disease after inoculation. Whereas S.
equi skin-test negative and seronegative horses (n = 8, group II) showed predictable
and severe clinical signs of infection after their inoculation, including shedding of the
pathogen from nasal discharge and ruptured submandibular lymph nodes. It was
concluded that resistance to virulent strain of S. equi infection is correlated with
existing humoral and cellular immune responses to Streptococcal antigens. The
horses that were prone, their recovery from infection were accompanied by the
appearance of antibodies and the positive skin-test response to S. equi antigens. Piche,
(1984) discussed the clinical observations during an epidemic of strangles that
occurred during the spring, summer and fall of 1980 on a Standardbred stud farm in
Eastern Alberta. S. equi might be introduced by a mare that was brought to the stud
form for breeding pupose. All of the horses on the farm were affected and for the
14
most part of the study that the disease was allowed to run its natural course. Only
badly affected horses were treated. During this outbreak, the foals were
prophylactically treated with penicillin to prevent them from contracting the disease.
Out of them 10 horses died due to complications of strangles infection. Timoney and
Trachman, (1985) studied the immunologically reactive proteins in acid extracts.
Then they adopt combination of chromatographic and immunologic procedures to
identify S.equi from culture supernatants. Both high and low molecular weight
components of each of these protein preparations were vigilant for mice. It was
surmised that there were variety of hydrolytic fragments of the M protein of S. equi
were present. Convalescent horse sera that exhibited strong bactericidal activity for S.
equi always reacted with polypeptides in the molecular weight range of 24,000 to
29,000, whereas pre infection sera did not. Rabbit anti sera to affinity-purified S. equi
protein also reacted with these polypeptides, as well as with a polypeptide of about
36,000 to 37,000 molecular weight. While M protein in acid extract and culture
supernatant did not cross-react in immunodiffusion, rabbit antiserum to affinity-
purified M protein from an acid extract of S. equi reacted aggressively with culture
supernatant proteins of approximate molecular weights of 67,000, 58,000, and
43,000. We suggest, therefore, that the M protein in culture supernatant is covered by
other sequences that are removed by hot acid during acid extracts preparation.
Evermann et al. (1987) find out the frequency of respiratory diseases in the horses in
North Western USA. The samples from clinically affected horses showed that both
equine influenza and S. equi were involved. Endemic strangles which occurred in all
horses that were susceptible, but mainly foals were affected showed pyrexia,
15
inflammation of the respiratory tract and lymphadenitis. Clabough, (1987)
investigated various aspects of strangles infection. He described 100% morbidity and
1-2% mortality. He concluded that mortality was due to dissemination of the infection
to other parts of the body. Once introduced, S. equi was a consistent problem due to
persistent ambient contamination. The infective organism was present in nasal and
abscess discharge from the infected horses. The IP of the disease was usually 4-8
days and there was a sudden onset of anorexia and fever. Initially Nasal discharge
was serous and developed within 24 hours, then it became purulent as the disease
attained its peak. Control of the disease was best accomplished by isolating the
affected horse and adopting strict hygienic measures by personnel handling the
animals. Gumbrell, (1987) reported that incidence rate of strangles had increased
significantly in Canterbury around Christchurch and normally 1-5 cases were
confirmed every month at the Lincoln Animal Health Laboratory. Mayr, (1987)
reviewed the major respiratory infectious diseases in horses and found that among
major diseases of horses, respiratory diseases pose the greatest threat to horses. The
most important respiratory infections of horses were caused by S. equi. Yelle, (1987)
published an article dealing with clinical aspects of S. equi. He concluded that the
incubation period of the disease and the course of the disease was 3-8 days and 3-4
weeks respectively the course of disease was usually 3-4 weeks reported. Morbidity
rate was too high 100% in susceptible population and the mortality rate was 2-3% if
the appropriate treatment was given. He described various risk factors and observed
that overcrowding and parasitism may increase the risk of the disease. Muhktar and
Timoney, (1988) observed the chemotactic response of leukocytes to S. equi infection
16
in horses. They concluded that the response was due to intense infiltration of lymph
node with polymorphonuclear leukocytes which suggested an influential chemotactic
response to the S. equi. Wilson, (1988) reviewed various aspects of strangles. He
found that S. equi is the causative organism and confirmed it as the major cause of
economic losses to the horse industry throughout the world. Because of the current
control measures were not completely effective in preventing the disease. The disease
affects only equines (horse, mules and donkeys). It was common in all horse-raising
areas especially young ones. Strangles was commonly seen in weanlings, yearlings
and young horses; very less in foals under one month and horses over 5 years of age.
This reflected very specific immunity, and horses of any age were susceptible if not
previously exposed or vaccinated. Overcrowding, movement and mixing of horses
from different sources are important predisposing factors. Sweeney et al. (1989)
studied the epizootic and infections of S. equi in horses. The age-specific attack rates
of S. equi infections in horses for the different age groups were calculated 17.6% for
broodmares, 47.5% for 1-year-old horses, and 37.5% for foals. S. equi was isolated
from different nasal swab, pharyngeal, or lymph node specimens in 31 (60.8%) of 51
disease horses. A male 1-year-old horse that had been taken from Kentucky to farm
A, was considered the index case. Six (19.4%) out of 31 horses with strangles
remained carriers for S. equi after clinical signs of the disease had ended. These
horses were not found positive for S.equi. After that they were kept with those horses
that were infected with S equi but strangles was not developed. Jorm, (1990) studied
Strangles in horse studs and calculated incidence, risk factors and effect of
vaccination. 179 horse studs in New South Wales was conducted to estimate the
17
incidence of strangles during that specified period 1985 to 1988, to identify risk
factors associated for strangles epidemics. During this particular period of time, forty-
nine studs (27.4%) had at least a single epidemic of strangles and 62 stud farms
(34.6%) had only a case of strangles. The average incidence rate of strangles was
calculated as 2.1 cases per 100 horses per year. The risk of strangles increased with
the increased horse population and rose markedly when more than 100 mares had
been served in the 1988-89 season. Different types of feeders, fences and water
sources were also significantly increased the chance of strangles outbreaks. Boyes et
al. (1991) reported a case of S.equi infection in which panophthalmitis also occurred.
The S. equi was isolated from corneal ulcer. The ulcer developed due to an extension
of septic uveitis subsequent to submandibular lymphadenopathy. The condition was
refractory to treatment and panophthalmitis ensued. S. equi was isolated from the
anterior chamber of infected eye through enucleation. It was observed that uveitis was
associated with S.equi, especially in those cases which had the history of strangles.
Hamlen et al. (1992) studied the hematologic parameters of 23 foals at weeks 0, 2, 4,
6, and 10 following the onset of a strangles epizootic. The epizootic was initiated by
group exposure with S, equi to a foal experimentally. The group was consisted of 12
foals, 6 months of exposure with S. equi epizootic, and 11 foals, previously
unexposed. It was observed that 91% of the unexposed and 17% of the previously
exposed foals developed clinical signs of strangles. Significantly increase in mean
WBC count, neutrophil cell count, fibrinogen concentration and plasma protein
concentration were seen in strangles cases as compared to foals not classified as cases
and were associated with clinical signs. Similarly, animals suffered from strangles
18
showed decreased PCV, Hg concentration, and RBC count, although statistically
insignificant, as compared to those without strangles during 4th, 6th, and 10th week that
may have biological significance. Similar, but more pronounced, changes in the
hematological parameters of the foal were observed in in those cases where S. equi
was inoculated experimentally. Besides of this effect of S. equi infection on the
hematology of foals should be considered in their convalescent care. Zadeh et al.
(1992), in Tehran, investigated the epizootology of strangles in equine. According to
an epidemiological survey 89-100% of horses were affected with the strangles. In
young horses the clinical signs were very severe but mortality rate was
approximately zero. All of the animals responded to these antibiotics
penicillin/streptomycin. Wood et al. (1993) studied the persistent infection with that
organism S. equi. They detected S. equi in the nasopharynx from one horse in the
severe outbreaks. They concluded that effective control measures of outbreaks of
strangles required monitoring of horses through swabbing. Dalgleish et al. (1993)
studied an outbreak of strangles infection in young ponies. A natural outbreak of
strangles occurred in 19 young experimental ponies. The disease was diagnosed in 11
of them within just two days of their arrival at Glasgow University veterinary school
and five others ponies were developed clinical signs within four days, a morbidity
rate reached upto 84%. All of the affected ponies had typical signs of strangles
disease including pyrexia, dullness, anorexia, regional lymphadenitis, occasionally
seen with rupture of the lymph node, conjunctivitis and mucopurulent nasal
discharge. 9 out of 19 ponies were destroyed during the clinical phase of the disease
for seeing post mortem changes. The clinical disease in the remaining animals lasted
19
upto 21 days although one pony had to be destroyed 10 days after the onset of clinical
finding because of the development of septic arthritis. All 16 affected animals
exhibited blood neutrophilia and high plasma fibrinogen levels. Beta haemolytic
streptococci were then isolated by nasopharyngeal swabbing from 18 out of 19
ponies. While the S. equi was confirmed in three animals within the first four days of
the outbreak. The majority of the other isolates identified to species were S
zooepidemicus. Beta haemolytic streptococci were still present in 6 ponies after 40
days and they had clinically recovered and were isolated on regular basis from these
three ponies which did not develop clinical strangles but remained in contact with
affected animals throughout the study. Timoney, (1993) illustrated the etiology;
epizootiology, pathogenesis, and clinical presentation of strangles. Streptococcus equi
is highly host specific to equine and shows no antigenic variation. Apparently
protective immunity is produced by a combination of serum opsonic and
nasopharyngeal mucosal antibodies responses. Vaccines based on M protein or
attenuated bacterial suspensions abate the clinical attack rate up to 50%, much lower
level of protection than that produced during recovery from strangles. Hamlen et al.
(1994) studied epidemiological and immunological features of S. equi infection in
different foals. A multiphase study was done to indicate the effects of S. equi
infection in previously exposed and unexposed foals. In phase I, they observed 22
weanling foals involved in a naturally occurring S equi epizootic, along with 11
unexposed foals for comparison, matched for sex, age, and breed. After six months
(phase II), an epizootic was experimentally induced in previously exposed and
unexposed foals from phase I and studied the prevalence, duration of clinical signs,
20
the relative risk of developing disease, bacteriologic culture results, hematologic
responses, mucosal and serum immunologic responses were determined. The
protection from disease in phase-I and -II foals was associated with high values for
serum S equi M protein-specific IgG at the onset of the epizootic (for phase 1 P <
0.02 and for phase II P < 0.01), and with a rapid (within 2 weeks of exposure)
mucosal S equi M protein-specific IgG response (for phase I P < 0.05 and P = for
phase II 0.01). Dwyer, (1995) compared the disinfectants of equine ambience that
provoke managers to take this challenge. For this purpose variety of surfaces which
may be contaminated with wide range of horse pathogens were used. The most
frequently occurring infectious diseases for which disinfection and disease control are
very important are strangles, salmonellosis and rotavirus diarrhoea. These are the
most difficult to control. The phenolic disinfectants have been demonstrated to be
effective in the presence of organic matter and are also have property of being
virucidal. When used after thorough cleaning and rinsing of stall surfaces, phenolic
disinfectants have proved effective in controlling outbreaks of different diseases.
Inspite of this, 10% iodophors used for hand washing and cleaning equipment is also
bactericidal and virucidal. Quaternary ammonium compounds like bleach,
chlorhexidine and pine oil are available commercially, but these disinfectants have no
consequence in the presence of the organic matter. Golland et al. (1995) carried out a
retrospective study on 46 horses suffering from retropharyngeal lymph node
infection. Horses aged <1year were observed oftenly affected (46%). A high
percentage of cases (39%) then exposed to horses and results were confirmed or
suspected strangles. The most frequent signs were unilateral or bilateral swelling of
21
throat region (65%), respiratory dyspnoea (38%), purulent nasal discharge (20%),
inappetence and signs of depression (15%) and dysphagia (9%). Along with
rhinopharyngoscopy, ultrasonography, haematology, cytological and microbial
analysis of material, aspirated from swelling of soft tissue, also helped in diagnosis of
the disease. Anzai et al. (1997) isolated the organism S. equi from thoroughbred
horses from racehorse-breeding area of Japan. During the breeding season in 1995 it
was observed whether strangles has spread in Hidaka district of Hokkaido and the
main racehorse-breeding area of Japan. For this aim an epizootiological survey with
bacterial isolation was conducted. S. equi was isolated from two Thoroughbred horses
showing signs of submandibular lymphadenitis. Then these isolates were
characterized and classified by serological grouping, different biochemical tests and
analysis of cell surface proteins by Western immunoblotting. From that survey, it is
end noted that S. equi has invaded the Hidaka district and in racehorse-breeding farms
in this area that strangles has become prevalent. Timoney et al. (1997) compared the
sequences and functions of, S. equi M-Like proteins, SeM and SzPSe. S. equi, a
Lancefield group C Streptococcus, causes strangles, contagious purulent
lymphadenitis and pharyngitis of members of the Equidae family. The antiphagocytic
M-like protein (58-kDa) SeM is a prominant virulence factor and protective antigen.
The amino acid sequence and structure of SeM has been known and equated to that of
a second, M-like protein SzPSe (40-kDa) of S. equi and along with other
Streptococcal proteins.It was shoen that SeM and SzPSe have no homology except
their signal and anchor sequences of the membrane and are alpha helical fibrillar
molecule. It was also observed that they have clear heterozygosity with other
22
Streptococcus M and M like proteins. Present study tells us that SzPSe is an allele of
SzP protein that encodes the mutable protective M-like and typing antigens of S.
zooepidemicus. Opsonogenicity of SeM only for S. equi. In contrast to SeM protein
the SzPSe is highly opsonogenic for S. zooepidemicus but not for S. equi. In the blood
SeM and SzPSe bind with equine fibrinogen. When they measured the size of SeM
and SzPSe of geographically and temporally separated isolates of S. equi they found
that there is no difference in size. This syudy shows that the S.equi is a clonal
pathogen of S. zooepidemicus. Al-Ghamdi et al. (2000) evaluated the use of repetitive
sequence-based PCR for molecular epidemiological analysis of S. equi. Inception of
the study was with 63 S. equi isolates from different areas of the world and 17 S. equi
isolates were collected during epidemics of S. equi. Then an aliquot of S. equi
genomic DNA was processed through PCR, using the specified enterobacterial
repetitive intergenic consensus primers. After this they run the samples on 1.5%
agarose gel and used software to equate these rep-PCR results. When they used these
primers to analyze 100ng genomic DNA of S. equi they observed the pattern of 6 to
14 bands. The initial isolates of 32 were segregated into 7 rep-PCR subtypes. They
also found 30 rep-PCR subtypes among 29 S equi isolates collected from Michigan,
Minnesota, Australia, Canada and 34 S equi isolates obtained from Kentucky State
and other sources when the epidemic of disease occurred, the same clone was
identified in several horses. All infected horses on the same farm had a single clone of
S equi. They concluded that this rep-PCR was most authentic for depicting S. equi
into various rep-PCR subtypes. Besides of this the results further disclosed that
isolates with the same geographic source and date of collection, did not have the same
23
rep-PCR subtype. A single clone of S equi usually predominated in an epidemic.
Chanter et al. (2000) examined S. equi with truncated M-proteins isolated from
outwardly healthy horses. The M-protein genes of S. equi isolated from 17 healthy
horses after four strangles outbreaks, including a quarantined animal, were compared
with S. equi isolates from 167 active cases of strangles across four countries. Sixteen
most occuring S. equi carriers were included in healthy horses, one from each of the
four outbreaks. These outwardly healthy carriers had empyema of the guttural pouch,
an increase in the size of equine Eustachian tube. A persistent carrier from two of
these outbreaks, the healthy animal and quarantined animal with normal guttural
pouches, from which S. equi was isolated only once, were colonized by variant S.
equi with truncated M-protein genes. The truncated M-protein genes had in-frame
deletions between the signal sequence and the central repeat region in slightly
different positions, equivalent to approximately 20% of the expressed protein. It was
end noted that immunoblotting with antibody to recombinant M-protein finalized that
the variants showed a truncated form of the M-protein. In contrast to the outwardly
healthy carriers of S. equi, only 1/167 of S. equi isolates from strangles cases
possessed a truncated M-protein gene. Comparing the isolates of healthy horses with
a truncated M-protein, much more of the N terminal of the truncated M-protein was
retained in the variant S. equi from a strangles case. Variant S. equi from outwardly
healthy horses were more susceptible to phagocytosis by neutrophils in vitro than
typical isolates. This was the first report of identification of truncated M-protein in S.
equi. The distribution of these variants between infected cases and carriers proposed
that the 80% of the M-protein retained in the variants that may contribute to
24
colonization whilst the deleted portion of the gene may be cause of full virulence.
Newton et al. (2000) studied the control of strangles outbreaks by isolation of guttural
pouch carriers through PCR and culture of S equi. From the previous study the use of
repeated nasopharyngeal swabbing and culture of Streptococcus equi visualized that
healthy carriers developed in more than 50% of strangles outbreaks. The guttural
pouches were the only site where S. equi colonisation on endoscopic examination of
horses during one of these outbreaks and sometimes S. equi was not identified by
culture of nasopharyngeal swabs for carriers up to 2 or 3 months before nasal
shedding resumed sporadically. Therefore more sensitive way of detecting S. equi on
swabs from established guttural pouch carriers was required. Strangles outbreaks
were reported in detail using endoscopy, in order to make development and
assessment of a suitable PCR test. From 3 protracted strangles outbreaks on different
kinds of establishment’s ranges between 29 and 52% of sampled horses were infected
as noticed by culture and PCR. Out of these, between 9 and 44% were identified as
carrying S. equi after clinical signs had abolished and the most predominant site of
carriage was the guttural pouch. Prolonged carrier of S. equi, which lasted up to 8
months, did not stop spontaneously before treatment was incepted to remove the
infections. For identification and separation of the carrier animals, along with strict
hygienic conditions, apparently resulted in the control of strangles outbreaks and
allowed the premises to come to normal activity. To compare PCR and culture, many
more swabs were found to be positive using Polymerase Chain Reaction (56 vs. 30%
of 61 swabs). Similarly for guttural pouch samples from 12 established carriers (PCR
76% and culture 59%). PCR can only identify dead organisms and is, thus, liable to
25
produce false positive results. Verheyen et al. (2000) studied to treat guttural pouch
infection and inflammation in asymptomatic carriers from this S. equi. Three
outbreaks of strangles were diagnosed by endoscopic study and a total of 14
asymptomatic carriers of S. equi were noticed out of which 13 showed evidence of
carrier state in the guttural pouch. Then treatment was started to eliminate S. equi.
Two other horses were referred to them with severe guttural pouch pathology and
from which S. equi was also cultured, and treatment of these cases was also
described. In the first instance treatment was directed towards removal of gross
guttural pouch pathology as seen on endoscopic examination. This was done by using
a combination of irrigation of the pouch with moderate to large amounts of saline
then suction of fluid and endoscopic manipulation of chondroids. Antibiotic treatment
was used to cure S. equi infection. Systemic antibiotics were given to all animals, but
in some cases along with antibiotics topical antimicrobial treatment was also given.
Treatment was generally considered as successful when the guttural pouches
appeared normal and S. equi was not noticed in nasopharangeal swabs and completed
the pouch lavages on 3 consecutive occasions. Successful treatment of one carrier
required surgical intervention due to occlusion of both guttural pouch pharyngeal
openings. 14 out of 15 carriers were successfully treated by endoscopic elimination of
inflammatory material and antibiotic treatment, without any surgical intervention. 5
carriers originally given potentiated sulphonamide (33%) required for further therapy
with combination of penicillin or ceftiofur, given systemically and topically, before
whole S. equi infection and associated inflammation of the guttural pouches were
completely removed. Harrington et al. (2002) studied Streptococcus equi is
26
responsible for strangles, one of the most dominated diseases of the equine. The
animal suffering and economic burden associated with this disease need effective
treatment. Current antibiotic treatment is often ineffective and thus recent attention
has focused on vaccine production. A systematic understanding of S. equi virulence,
leading to the identification of targets to which protective immunity can be directed,
is a prerequisite of the development of such a vaccine. Here, the virulence factors of
S. equi are studied. Ensink et al. (2003) evaluated the efficacy of
trimethoprim/sulfadiazine and procaine penicillin G against S. equi subsp.
zooepidemicus in ponies. Tissue chambers, implanted s/c on both sides of the neck,
were inoculated with S. zooepidemicus to compare the efficacy in a purulent
infection. The TMP/SDZ treatment consisted of one i.v. injection of 5 mg/kg TMP
and 25 mg/kg SDZ and the same dose of TMP/SDZ per os both given 20 h after
inoculation. Then the oral dose was repeated every 12 h for 21 days. The penicillin
treatment was given i.v. injection of 20 000 IU/kg sodium penicillin G and i.m.
injection of 20 000 IU/kg procaine penicillin G, both given 20 h after infection. Then
the i.m. dose was repeated after every 24 h for 21 days. TMP/SDZ resulted in a
limited reduction of viable bacterial count in the TC but did not remove the infection,
thus resulting in abscess formation in 10-42 days in all eight ponies.While, penicillin
eliminated the streptococci in 7 of 8 ponies, and only one pony suffered from abscess
formation on day 10. It is concluded that penicillin showed significantly better
efficacy than TMP/SDZ. Therefore, it is recommended that TMP/SDZ should not be
used to treat purulent infections in secluded sites in horses. Masakazu et al. (2003)
observed fever and enlarged submandibular lymph nodes were seen during quarantine
27
inspection in 3 out of 9 quarter horses imported from the USA. They were found to be
negative in bacterial examinations during the quarantine period then these horses
were released. Riding club was their destination; however, S. equi was isolated from 3
of them. Then strangles infection spread to other horses in the riding club. Nasal
swabs were obtained from all horses at the club on weekly basis until the disease
subsided 22 weeks later. S. equi was isolated from 25 of 58 horses (43.1%). It was
concluded that the imported carrier horses were responsible for the spreading of
infection throughout the riding club. Dwyer, (2004) compared the different
environmental disinfectants to control equine infectious diseases. Therefore it is
recommended that cleaning and disinfection is essential to the environmental control
of infectious diseases of all animals. To see the types of pathogens, environment,
disinfection process and success can be attained in effectively by stopping disease
outbreaks. Timoney, (2004) studied the important and potent pathogenic Streptococci
for horse which includes S. equi, S. zooepidemicus, S. dysgalactiae subsp. equisimilis
and S. pneumoniae capsule Type III. S. zooepidemicus strain is the ancestor of S.equi,
both shares more than 98% DNA homology and that is why expresses many common
proteins and virulence factors. Strategic progress has been made for identification of
different virulence factors and proteins which are only expressed by S. equi. Most of
these proteins and virulence factors are expressed on bacterial surface. The notable
examples include antiphagocytic SeM, the secreted pyrogenic superantigens SePE-I
and H. The genomic DNA sequence of S. equi will greatly help in identification and
characterization of more virulence factors and vaccine targets. S. equi is the most
frequently isolated opportunist organism of the horse. Gronbaek et al. (2006)
28
evaluated a nested PCR test and bacterial culture from nasal swabs and abscesses for
the diagnosis of S. equi infection (strangles). When he used nested PCR all 45 S.equi
were positive and not a single amplicon was formed when testing the other 120
Lancefield group C isolates. He collected 43 samples from 11 horses that were
showing typical clinical signs of strangles. He investigated the diagnostic sensitivity
for PCR test was 45% and 80% for samples from the nasal passages and abscesses,
respectively; whereas diagnostic sensitivity for cultivation was 18% and 20%.
Therefore it is stated that diagnostic sensitivity was significantly higher for PCR than
for bacterial cultivation. Along with this, they evaluated the shedding of S. equi in 2
infected horse populations. They observed the intermittent shedding period of S.equi
which was up to 15 days. Shedding of S. equi was seen in both from horses with and
without clinical signs. They finally concluded that nested PCR test is highly species-
specific and is very sensitive method for detection of S. equi from clinical samples.
Kelly et al. (2006) observed the sequence variation of the SeM gene of S. equi allows
discrimination of the source of Strangles outbreaks. Improved understanding
regarding, the epidemiology of S. equi transmission demands high sensitivity and sub
typing methods that can rationally differentiate between strains. S. equi is highly
homogeneous and cannot be distinguished by multilocus sequence-typing or
multilocus enzyme electrophoretic methods that use housekeeping genes.
Nevertheless, sequential analysis of the N-terminal region of the SeM genes of 60 S.
equi isolates from 27 outbreaks of strangles, find out 21 DNA codon changes. These
resulted in the non synonymous replacement of 18 amino acids and allowed the
assignment of S. equi strains to 15 distinct subtypes. The findings of the present study
29
propose the presence of multiple epitopes across this region that is subjected to
selective immune pressure, especially in the establishment of chronic S. equi
infection. They further pointed out the the application of SeM gene sub typing
procedure to examine potential cases related to administration of a live attenuated S.
equi vaccine. SeM gene sub typing discriminated between the vaccinal strain and field
strains of S. equi the actual cause of disease. The results were confirmed by the
establishment and application of a PCR test, which identifies the aroA partial gene
deletion present in the Equilis StrepE vaccine strain. It was concluded that the
injection site lesions were due to vaccinal strain, all seven outbreaks of strangles
investigated in recently vaccinated horses were found to be due to agreeing infection
with wild-type S. equi and not reversion of the vaccine strain. Tiwari et al. (2007)
examined Se18.9, an anti-phagocytic factor H binding protein of S. equi. The escape
from phagocytosis is of great significance in virulence determination of S. equi, the
cause of strangles in equine and discriminate it from the closely related S.
zooepidemicus. They explained Se18.9, a novel H factor binding protein which is
only secreted by S. equi and not by S. zooepidemicus that reduces accumulation of C3
on the surface of bacteria and in consequence reduces the bactericidal activity of
neutrophils significantly suspended in normal serum for both S. equi and S.
zooepidemicus. Se18.9 is produced and released in large quantity by actively dividing
cells abundantly and is also bound to the surface of bacteria. Strong mucosal and
serum antibody responses are elicited in S. equi infected horses. Although there was
no gene identical to Se18.9 in S. zooepidemicus, sequences encoding proteins of same
size with closely related signal peptide sequences were found in 3 of 12 randomly
30
selected strains. As Se18.9 is specific to S. equi, and immunoreactive with mucosal
IgA and convalescent sera, so it could be used for diagnostic purpose. Waller and
Jolley, (2007) investigated getting a grip on strangles. Strangles, that remained one of
the most commonly diagnosed and a high ranked infectious disease of horses through
world-wide. This review article elaborated the diagnosis and pathogenesis of
strangles and is fastidious to the importance of prolonged infection with particular
attention to the significance of persistent infections. Now it is possible to combine
recent sequence data obtained from the N-terminal site of the SeM to reallocate the
SeM alleles with the help of on-line database. Hypotheses concerning the inception of
this variation and the capability of being exploit for the epidemiological investigation
of outbreaks are suggested. Zadeh et al. (2007) concluded that strangles is an acute
disease of horses caused by S. equi and characterized by inflammation of the URT
and abscess formation in the lymph nodes, distributed worldwide. Outbreak occurs
mostly in a large population of young horses and affected young ones upto 100%.
High incidence is observed soon after the gathering of large number of susceptible
horses, which may have come from different areas and are stabled together. The
infection source is the nasal discharge from infected animals, which contaminates the
pasture, feed and water troughs. Infection occurs either by ingestion or by inhalation
of droplets. Two cases of strangles were diagnosed at Veterinary Faculty Teaching
Hospital in Tehran during April (1990). Then an epizootological survey immediately
started and showed 89-100% of horses were in incubation stage. The clinical signs
were intense in younger horses but the mortality rate was zero. All of the infected
horses gave good response to the treatment with i/m injection of
31
penicillin/streptomycin. Jannatabadi et al. (2008) checked the existence of S. equi as
possible causative agent of upper respiratory tract infection in Mashhad area. Nasal
swabs samples were collected from 30 horses with URT infections. Then different
types of bacteria were isolated from samples S. equi (1 isolate), S. zooepidemicus (25
isolates) and others. Confirmation of these isolates of S. equi and S. zooepidemicus
were done by different biochemical tests and Polymerase chain reaction. For further
molecular identification of S. equi and S. zooepidemicus, two genomic region SeM
and sodA were amplified. Knowles et al. (2010) tested 30 horses having no external
clinical signs of strangles for exposure to S. equi using a latest serological test. Inspite
of this, carrier state of S. equi was also checked by using endoscopy of the guttural
pouches and PCR. Serological results were showed that four horses had been recently
exposed to S. equi and started non-specific clinical signs of respiratory disease. One
asymptomatic horse out of four was also positive for S. equi by PCR.
32
Chapter-03
MATERIALS AND METHODS
The present study was conducted on strangles in equines of Lahore and
Sargodha districts of the Punjab province of Pakistan. The samples collected were
processed at Medicine and Microbiology Laboratories of the University of Veterinary
and Animal Sciences, Lahore. Pakistan and Gluck equine research center, Department
of Veterinary Science, University of Kentucky, USA. The study was comprised of
five phases as under.
PHASE I:
Epidemiology of Disease:
In this phase of epidemiology, prevalence of the disease, variations in SeM,
SzPSe, Se18.9 proteins and mortality rate were studied. For prevalence a total of 500
equines (n=250 horses; n=250 mules) was examined from Lahore and Sargodha
districts of Punjab province. The data related to equines were collected in a data
capture form. The entries in data capture form included location, species, age, sex,
breed, season, previous disease history and mortality.
Prevalence:
Prevalence was referred to the amount of disease in each district for a period
of one year, without distinction between old and new cases were calculated as per
formula described by Thrusfield, (2002).
No. of individual having a disease at a particular point in time P = --- ------------------------------------------------------------------------------
No. of individuals in the population at risk at that point in time
33
Mortality Rate
The measure of the number of deaths in each district was calculated as per
procedure described by Thrusfield, (2002) as given below.
No. of deaths due to disease that occur in a Population during a particular period of time
Mortality Rate = ------------------------------------------------------------- The sum, overall individuals, of the length of time at risk of developing disease
Collection of Samples:
Following two types of samples were collected aseptically for identification of
causative agent (S. equi) of strangles.
a) Nasal discharge was collected from nasal chambers using sterile cotton swabs.
b) Pus from affected lymph nodes was collected aseptically using sterile
disposable syringes (Merchant and Packer, 1983).
The collected samples were processed at Medicine & Microbiology
Laboratories, University of Veterinary and Animal Sciences (UVAS), Lahore
Pakistan and Gluck equine research center, Department of Veterinary Science,
University of Kentucky, USA.
Culture and isolation of S. equi:
The samples were cultured on blood agar plates and incubated anaerobically
for 24 hours to optimize the isolation of the organism preferentially against the other
organisms present in the nasal passages (Jorm, 1990).
Confirmation of S. equi by using biochemical test:
The above isolates were identified on the basis of cultural, morphological and
biochemical characteristics following the techniques as described by Buxton and
Fraser (1975) and Merchant and Packer (1983).The biochemical tests to be carried
34
out were include, Catalase reaction, Methyle blue reduction and Sugar fermentation
test ( Trehalose, Latose, Manitol, Salicin and Maltose). Isolates identified as S. equi
fermented salicin and sucrose, but not lactose, sorbitol, trehalose. (Quinn et al.,
1994).
Colony characteristics of S.equi Beta hemolytic Pattern of S. equi
Selection of beta haemolytic colonies:
Pure β-hemolytic colonies on blood agar were selected for DNA extraction.
Genomic DNA purification kit method:
1. Take 10-20mg of bacterial (S. equi) culture and resuspend in 1.5ml eppendorf
containing 200µl TE buffer.
2. Add 400µl of Lysis solution and mix it.
3. Incubate at 65ºC for 5 minutes.
4. Add 600µl Chloroform and gently emulsify by inversion.
5. Centrifuge at 10,000 rpm for 2 minutes.
6. Transfer the upper aqueous phase containing DNA to a fresh tube.
35
7. Add 800µl precipitation solution (To prepare fresh working precipitation
solution, mix 720µl nuclease free water with 80µl of supplied 10X
concentrated precipitation solution.).
8. Mix at room temperature for 1-2 minutes.
9. Centrifuge at 10,000 rpm for 2 minutes.
10. Remove the supernatant completely.
11. Dissolve the DNA pellet in 100µl of 1.2M NaCl solution. Make sure that the
pellet is completely dissolved.
12. Add 300µl cold absolute ethanol and let the DNA precipitated (10 minutes at -
20ºC). This time can be extended upto overnight stay at -20ºC.
13. Centrifuge at 10,000 rpm for 6-8 minutes.
14. Discard the supernatant and keep the pellet.
15. Wash the pellet once with 100µl 70% cold ethanol.
16. Let the pellet dry completely.
17. Dissolve the DNA pellet in 100µl of autoclaved double distilled deionized
water.
Quantification of Extracted Genomic DNA:
Quantification of extracted DNA was done using 0.8% Agarose gel. A
standard DNA ladder (50bp) was run with the sample for quantification. Picture was
taken by gel documentation system for record. All samples were brought to same
concentration.
36
POLYMERASE CHAIN REACTION:
Primer Selection and Optimization:
Primers for SeM gene of S. equi were showed in table 3.1. These primers
were optimized by gradient PCR using thermocycler. The PCR product was run on
1.2% Agarose Gel.
PCR amplification:
PCR amplification was carried out using S. equi specific primers. These
primers amplify 677bp and 325bp region of SeM gene of S. equi. The PCR was
carried out with the reaction mixture of 25 μl and the quantities of other components
are as under.
2x Taq PCR buffer 11 µl
ddH2O 07 µl
Primer F (3µM) 2.5 µl
Primer F (3µM) 2.5 µl
Sample DNA 02 µl
Total 25 µl
The PCR was carried out in four steps. 1st step was initial denaturation at
950C for 4 min then for each of 30 cycles, the denaturation at 950C for 1min,
annealing for 1 min and 3rd stage extension at 720C for 2 min. PCR cycles were
followed by 10min of final extension at 720C. And at the end, final holding
temperature was 40C until the PCR tubes were taken out of thermal cycler and placed
in refrigerator or run on Agarose Gel.
37
Table 3.1: Details of all the primers used in the study.
4. Analysis of PCR products (Agarose gel electrophoresis):
1. The amplified PCR products were analyzed by electrophoresis at 80V and
300mA in 1.2 % agarose gel for one hour and thirty minutes using 1X TAE
buffer.
2. Desired amount of agarose (1.2g) was taken in flask containing the
electrophoresis buffer 1X TAE (100ml), melted in microwave oven for 2
minutes and swirled to ensure even mixing.
Primer Sequence Amplicon
(bp)
Annealing
Temp. Source
SeMF
SeMR
TGCATAAAGAAGTTCCTGTC
GATTCGGTAAGAGCTTGACG 677 560C
Jannatabadi et al., 2008
SeMF
SeMR
CATACCTATCTCCATCAGCA
CGAACTCTGAGGTTAGTCGT 325 570C
Jannatabadi et al., 2008)
IGSzMF
IGSzMR
AAA GTG TGC CCA TAA CGG GTA
CGG CTA TTG TCC ATT GGG GAA 1812 650C
Present study
IGSZPF IGSZPR
CTT GCT AAA GTA ATG GTT GAC
GTT TGT GAG CAA GGC TTA GTC 1212 500C
Present study
SzPIPF SzPIPR
ATG GCA AAA AAA GAA ATG AAG
TTA GTT TTC TTT GCG TCT TGT TGA 1152 590C
Present study
IG18.7F
IG18.7R
ATA ACC ACT CGT TTA CAT GAG TG
GGT CCA AAA ATA CTA TTC TGA ACC 735 600C
Present study
Se18.7F
Se18.7R
AGT TTT AGC CAG TGC AGC AGC
TTA ATT CTC CAG ACT TTT CAA G 481 550C
Present study
EqbE -F
EqbE-R
AAG ATA TAG CAG CAT CGT ATC G
TCT AAA TCT CTA TTA AAT AGC GGT ATA TTG
130 550C Heather et al., 2008
38
3. Casting tray was prepared by wrapping meshing tape on its both sides and a
suitable comb was adjusted on it for making the wells
4. The melted agarose was cooled to 45°C and Ethidium Bromide (5μl) was
added into it, before pouring on the gel casting tray.
5. The gel thickness was kept in the range of 0.5-1.0cm. The air bubbles were
removed and it was kept at room temperature for solidification.
6. After solidification the comb was removed carefully to avoid tearing of wells.
The mesh tape was also removed from the sides of the casting tray.
7. The gel casting tray containing the gel was placed in the electrophoresis tank,
having 1X TAE buffer.
8. PCR products (10-15μl) were loaded on the gel in the respective wells after
mixing with appropriate amount (4μl) of Invitrogen’s Blue Juice (6X gel
loading buffer).
9. 100 base pair DNA marker was run along with, as reference.
10. Electric current (80V, 300mA) was applied until the blue dye was about the
three quarters the way down the gel (approximately 90 minutes).
11. Gel was taken out of the electrophoresis tank when the dye had reached near
the anode part of the gel.
12. Gel was then photographed under the UV transilluminator at a wavelength of
254nm with Eagle Eye Gel Documentation System (Stratagene, USA).
Variations in SeM, SzPSe and Se 18.9 proteins of S.equi:
Twenty-five isolates of S. equi including CF32 (S. equi) as reference strain
from outbreaks of equine strangles in N. America, Europe, Australia and Pakistan
39
during the years 1969 to 2009 were selected for the study (Table 3.2). These included
pairs of sequential isolates from horse JB with a chronic infection of the maxillary
sinus and one horse 836 that experienced prolonged shedding. The collected samples
were processed in the Gluck equine research center, Department of Veterinary
Science, University of Kentucky, USA. All isolates of S. equi were negative for acid
production from lactose, sorbitol and trehalose. Each was propagated from single
colonies in Todd-Hewitt broth (THB) for DNA isolation. DNA from Pakistani
isolates S24 and L32 were extracted using a Genomic Purification Kit (Fermentas #
K0512). DNAs from the remaining S. equi isolates were obtained as follows. Pellet
from overnight culture in 10 ml THB was suspended in 150 μl digestion buffer (10
mM Tris HCl, 1 mM EDTA, pH 8.0) to which was added 10 μl lysozyme solution (5
mg/ml digestion buffer). Following incubation at 37ºC for 30 min, 20 μg proteinase K
(Sigma) in 5 μl digestion buffer was added followed by heating at 56ºC for 30 min.
Finally each bacterial suspension was boiled for 10 min, centrifuged at 10,000 x g for
5 min and the supernatant stored at -20ºC. Primers designed for amplification and
sequencing of SeM, SzPSe, and Se18.9 are listed in Table 3.1. The PCR protocol
consisted of 30 cycles of 94ºC for 1 min, annealing for 1 min, and 72ºC for 2 min.
PCR products were purified using GeneJET PCR purification kit (Fermentas) and
sequences (Eurofins MWG Operon) obtained using primers from the initial
amplification.
40
Table 3.2: Geographic origin, year of isolation and strain characteristics of S.equi .
Strai
n
Year
isolated
Geographic
Origin Strain Characteristics/Comments
S24 2010 Pakistan Isolated S.equi from mandibular abscess
L32 2009 Pakistan Isolated S.equi from nasal wash
MN08 1996 USA(MN) Mucoid S.equi from mild case of strangles
119 1191 Scotland Mucoid S.equi from typical strangles case.
Jba 1998 USA (NY) Mucoid S.equi from persistently infected ethmoid sinus
JBb 1996 USA (KY) Mucoid S.equi from persistently infected ethmoid sinus
FF 1998 USA (KY) Mucoid S.equi
836a 1997 USA (KY) Mucoid S.equi isolated from pony 836,experimentally infected with S.equi CF32
836b 1998 USA (KY) Mucoid S.equi isolated from pony experimentally infected with S.equi CF32
LEX 1990 USA (KY) Non mucoid S.equi from mandibular abscess
E33 1976 USA (NY) Non mucoid S.equi
181 1990 AUSTRALIA Mucoid S.equi from lung abscess 331 1982 Swedish Mucoid S.equi from typical strangles.
BM 1983 Ireland Matte colony of S.equi from case of strangles
M4 1991 Germany Mucoid S.equi
E9 1975 USA (NY) Matte S.equi from strangles case (nasal wash)
FT 2003 USA (IA) Mucoid S.equi from horse with post strangles myopathy
JP 2010 Denmark Mucoid S.equi from guttural pouch
NM 2007 USA (KY) Non mucoid S.equi
MER7
2007 USA (VA) Non mucoid S.equi
15C 2001 USA (UT) Mucoid S.equi from strangles case in captured wild horse
Lou 2010 USA (ME) Non mucoid raised colony of S.equi from mandibular abscess
107 1991 Scotland Mucoid S.equi from nasal wash from typical strangles
K58 1969 USA (KY) Mucoid S.equi from case of strangles
W60 1976 USA (NY) Non mucoid, prototype US strain
CF32 1981 USA (NY) Mucoid, prototype US strain
41
PHASE II:
Study of carrier animals:
A total of 40 equines (n=20 horses; n=20 mules) affected with strangles and
showed clinical sign of lymph node enlargement that remained positive after one
week of infection were monitored for 12 weeks to study the carrier status. Samples
were collected on weekly basis from these equines and were processed by using
culture and Polymerase Chain Reaction.
PHASE III:
Haematological Examination:
For haematological examination, 40 horses (n=20 healthy horses; n= 20
diseased horses) and 40 mules (n=20 healthy mules; n= 20 diseased mules) under 5
years of age suffering from natural outbreak of strangles and similarly for carrier
animals 40 horses (n=20 healthy horses; n= 20 carrier horses) and 40 mules (n=20
healthy mules; n= 20 carrier mules) recovered from strangles from Lahore and
Sargodha districts of Punjab province were included. The blood samples were
collected at weekly intervals from affected equines and monthly intervals form carrier
animals, directly from jugular vein, in sterilized plastic bottles coated with EDTA @
1mg/ml of blood. Following haematological parameters, total white blood cell count,
mean segmented neutrophils count, total lymphocytic count, total monocytic count,
total eosinophilic count, total basophilic count, total erythrocytic count, packed cell
volume and haemoglobin concentration were studied.
42
Protein Analysis:
For protein analysis 40 equids (n=20 horses, n= 20 mules) under 5 years of
age suffering from natural outbreak of strangles from Lahore and Sargodha districts
of Punjab province were included. Blood samples were collected from Jugular vein in
clean dried centrifugal tube without anticoagulant and brought to laboratory for the
separation of serum. Coagulated blood was centrifuged at 3000 rpm for 20 minutes
and with the help of pipette serum was transferred into clean dried bottles. Samples
were stored 40C for further analysis (Weichselbaum, 1946). Following parameters,
total serum protein, serum albumin, serum globulin and fibrinogen were studied by
using serum chemistry analyzer.
PHASE IV:
THERAPEUTIC TRIALS:
In-vitro testing of Antibiotics:
In this phase, in-vitro antibiotic sensitivity of S. equi to various antibiotics
(Procaine penicillin, ceftiofur Na, cephradine, erythromycin, ampicillin, tetracycline,
chloramphenicol, sulfamethoxazole, trimethoprim+sulfadiazine and gentamycin.) was
determined.
Procedure of Kirby-Bauer Antibiotic Sensitivity Test:
1. Inoculate all blood agar plates with S. equi.
2. Using the swab, streak the entire agar surface horizontally, vertically and
around the outer edge of plate to ensure heavy growth over the entire surface.
3. Allow the culture plates to dry for about 5 minutes.
43
4. Using the Sensi-disc dispenser, apply the antibiotic discs by placing the
dispenser over the blood agar surface.
5. Then gently pressed each disc so that they adhere to the surface of blood agar.
6. Incubated all cultured plates for 24 hours at 370C.
7. Measured zone of inhibition of each disc.
In-vivo Antibiotic Trials:
Based on the above in-vitro sensitivity test four top ranking antibiotics were
selected and administered to diseased animals and their efficacy were studied. Detail
of groups is as follows:
Groups No of animals
Antibiotic Horse Mules
A 10 10 1
B 10 10 2
C 10 10 3
D 10 10 4
The efficacy of antibiotics was checked on the basis of disappearance of
clinical signs.
PHASE V:
In-vitro Disinfectant Trial:
In vitro testing of S. equi with disinfectants to determine susceptibility of
bacteria to the disinfectants was performed. The disinfectants used in the study were
Dettol, Povidone iodine, 0.6% H2SO4 and Bleach. The efficacy of disinfectants was
compared by using the Phenol Co-efficient Test (Cappuccino and Sherman 2004).
44
Procedure of Phenol Co-efficient Test:
1- 45 nutrient broth tubes were labelled with the names and dilution of the
disinfectant and time interval of subculturing.
2- In test tube rack, six serial dilution 1:50, 1:100, 1:150, 1:150, 1:200, 1:250
and 1:300 of each disinfectant used in the present study (phenol, dettol,
povidone iodine, 0.6% H2SO4 and bleach) were made in normal saline in
separate tubes.
3- Rapidly inoculated one drop of the S. equi culture into each tube of
disinfectants and noted the time of inoculation.
4- All the test tubes were agitated to ensure contact between the disinfectant
and the microbes.
5- Using sterile technique, at intervals of 5, 10 and 15 minutes one loopful
was transferred from each of the test tubes into the appropriately labeled
sterile tube of nutrient broth.
6- All tubes were incubated for 48 hours at 370C.
The phenol coefficient was then calculated by using following formula.
Highest dilution of disinfectant that kill microorganism in 10 min. not in 5 min. -------------------------------------------------------------------------------------------------------
Highest dilution of phenol that kill microorganism in 10 min. not in 5 min.
Statistical Analysis
Data on prevalence of the diseases on the basis of culture and PCR were
analyzed by Chi-square test using statistical software package STATA 9.1 college
station T×77845, USA while data on haematological and biochemical examination
45
were analyzed with one way ANOVA. The difference between diseased and healthy
animal along different weeks was tested by Tukey’s test. A P-value <0.05 was used to
reject the null hypothesis that the model is not significant. Whereas sequences of
genes were compared with S. equi CF32 and with the database using multiple
sequence alignment by CLUSTALW. SeM alleles were identified using S. equi SeM
database www.pubmlst.org/szooepidemicus/ using BLAST. New unlisted alleles were
assigned numbers 71 to 76. Frequencies of single nucleotide polymorphism (SNP) in
SeM, SzPSe and Se18.9 were obtained by aligning DNA sequences of each isolate and
counting SNPs relative to those sequences in S. equi CF32. Inserted and deleted
sequences were not included in computation of SNPs. G-C percentages were obtained
using G-C Calculator.
46
Chapter- 04
RESULTS
To study strangles in equines present project was conducted in Lahore and
Sargodha districts of the Punjab province of Pakistan. The samples collected were
processed at the Medicine and Microbiology Laboratories of the University of
Veterinary and Animal Sciences, Lahore and Gluck Equine Research Centre,
Department of Veterinary Science, University of Kentucky, USA. Data on various
parameters were collected and analysed. Results are given below.
EPIDEMIOLOGY OF STRANGLES:
To study overall prevalence of strangles in equines a total of 500 equines
(n=250 horses; n=250 mules) were examined. On the basis of culture 113 horses and
99 mules were found positive for strangles. An overall prevalence of strangles was
recorded as 42.4% whereas prevalence of strangles in horses and mules was recorded
as 45.2% and 39.6 respectively. Prevalence once evaluated through PCR 122 horses
and 113 mules tested positive for strangles resulting in overall prevalence rate as
47%. Prevalence rate for horses and mules remained 48.8% and 45.2% respectively
shown in fig.4.1. It is obvious from the results that prevalence rate in both the species
in response to PCR were higher than those in response to culture.
47
Fig 4.1: Prevalence of Strangles in horses and mules
PREVALENCE OF STRANGLES IN HORSES:
Prevalence of Strangles in horses on the basis of culture:
The results of prevalence of strangles in nasal discharge and pus from sub-
mandibular lymph nodes of horses in different age groups and in different months of
the year are shown in table 4.1. The prevalence rate in horses under 1 year of age was
45(90%), between 1-2 years of age was 42(84%), between 2-3 years of age was
14(28%), between 3-4 years of age 8(16%) and 4-5 years of age was 4(8%). The
highest prevalence was recorded in the group having less than 1 year of age and then
between 1-2 years and decreased with increase in age. Prevalence in horses of
different age groups was observed to be significantly different from each other
(p<0.05). It can be concluded from the results that the horses under 2 years of age are
the most susceptible to strangles infection.
48
Table 4.1: Prevalence of Strangles in nasal discharge and pus samples of sub-mandibular lymph nodes of horses on the basis of culture.
Chi-square analysis showed significant difference in prevalence of strangles among all age groups (Chi-square value 122.02, P- value 0.0001)
Similarly, when the prevalence of strangles in horses during different months
of year was calculated, it was found to be the highest during the months of February,
March, April and May as shown in fig. 4.2. While few cases were seen during the
months of January, June and July and no cases were seen during other months of the
year.
Fig 4.2: Month wise prevalence of Strangles in horses on the basis of culture.
Age Groups
n Number positive for S. Equi Total
(%) Jan Feb Mar Apr May Jun July Aug Sep Oct Nov Dec <1year 50 02 09 11 13 07 02 01 00 00 00 00 00 45(90)
1-2year 50 01 08 15 12 02 04 00 00 00 00 00 00 42(84)
2-3year 50 00 03 05 03 03 00 00 00 00 00 00 00 14(28)
3-4year 50 00 01 02 03 02 00 00 00 00 00 00 00 08(16)
4-5year 50 00 01 00 02 01 00 00 00 00 00 00 00 04(08)
Total 250 03 22 33 33 15 06 01 00 00 00 00 00 113(45.2)
49
A significant difference was observed (p<0.05) among prevalence rates during
different months of the year. It was concluded from the results reported here, that the
strangles season, in Pakistan lasts from January through the month of July. Similarly
when compared the prevalence of strangles in different seasons of Pakistan. The
highest prevalence rates were recorded during the spring months. The reason might
be the maximum exposure and movement of horses during the months of spring as
equestrian activities are at the peak during these months.
Prevalence of Strangles in nasal discharges:
The prevalence of strangles in nasal discharges of horses on the basis of
culture in different age groups and in different months of year is shown in table 4.2.
The prevalence rate in less than 1 year of age was 29(87.88%), 1-2 years of age was
30(81.08%), 2-3 years of age was 6(17.14%), 3-4 years of age 6(13.33%) and 4-5
years of age was 3(6.82%) on the basis of culture. The highest prevalence rates in
nasal discharges of horses were recorded between 1-2 years of age and then under 1
year of age while the lowest prevalence rate was recorded in 4-5years of age.
50
Table 4.2: Prevalence of Strangles in nasal discharge of horses on the basis of culture.
Age
Groups n
Number positive for S. equi Total (%) Jan Feb Mar Apr May Jun July Aug Sep Oct Nov Dec
<1year 33 01 04 08 09 04 02 01 00 00 00 00 00 29(87.88)
1-2year 37 01 04 11 09 01 04 00 00 00 00 00 00 30(81.08)
2-3year 35 00 00 03 01 02 00 00 00 00 00 00 00 06(17.14)
3-4year 45 00 01 01 02 02 00 00 00 00 00 00 00 06(13.33)
4-5year 44 00 01 00 01 01 00 00 00 00 00 00 00 03(06.82)
Total 194 02 10 23 22 10 06 01 00 00 00 00 00 74(38.14)
Chi-square analysis showed significant difference in prevalence of strangles among all age groups (Chi-square value 100.09, P- value 0.0001) Prevalence of Strangles in pus samples of sub-mandibular lymph nodes:
The prevalence of strangles in pus samples of sub-mandibular lymph nodes of
horses on the basis of culture in different age groups and in different months of year
was recorded and is shown in table 4.3. The prevalence rate on the basis of culture in
horses under 1 year of age was 16(94.12%), 1-2 years of age 12(92.31%), 2-3 years
of age 8(53.33%), 3-4 years of age 2(40.00%) and 4-5 years of age 1(16.67%). The
highest prevalence rates were recorded in pus samples of sub-mandibular lymph
nodes of horses in less than 1 year of age then 1-2 years of age while the lowest
prevalence rate was calculated in 4-5years of age.
When comparing the prevalence rate of strangles in nasal discharges and pus
samples of sub-mandibular lymph nodes in different age groups of horses, it was
observed that number of strangles cases were significantly higher (p<0.05) in pus
samples from sub-mandibular lymph nodes as compared to nasal discharges.
51
Table 4.3: Prevalence of Strangles in pus samples of sub-mandibular lymph node of horses on the basis of culture.
Age
Groups n
Number positive for S. equi Total (%) Jan Feb Mar Apr May Jun July Aug Sep Oct Nov Dec
<1year 17 01 05 03 04 03 00 00 00 00 00 00 00 16(94.12)
1-2year 13 00 04 04 03 01 00 00 00 00 00 00 00 12(92.31)
2-3year 15 00 03 02 02 01 00 00 00 00 00 00 00 08(53.33)
3-4year 05 00 00 01 01 00 00 00 00 00 00 00 00 02(40.00)
4-5year 06 00 00 00 01 00 00 00 00 00 00 00 00 01(16.67)
Total 56 01 12 10 11 05 00 00 00 00 00 00 00 39(69.64)
Chi-square analysis showed significant difference in prevalence of strangles among all age groups (Chi-square value 19.90, P- value 0.0001) Prevalence of Strangles on the basis of Polymerase Chain Reaction:
Likewise, the prevalence of strangles on the basis of Polymerase Chain
Reaction in different age groups of horses and in different months of year is shown in
table 4.4. The prevalence rate under 1 year of age was 46(92%), 1-2 years of age was
43(86%), 2-3 years of age was 17(34%), 3-4 years of age 10(20%) and 4-5 years of
age was 6(12%). The highest prevalence was recorded in the group of horses having
less than 1 year of age then 1-2 years and decreased with increase in age. The
significant differences were observed (p<0.05) among horses of different age groups.
Therefore it is concluded from the present study that the sensitivity of PCR appears to
be much greater than culture for nasal and pus samples from affected sub-mandibular
lymph nodes. As PCR can be completed in 4-5 hours hence is considered to be an
effective tool in diagnosis, control and management of strangles infection.
52
Table 4.4: Prevalence of Strangles in nasal discharge and pus samples of sub-mandibular lymph nodes of horses on the basis of PCR.
Chi-square analysis showed significant difference in prevalence of strangles among all age groups (Chi-square value 113.12, P- value 0.0001)
Similarly the year round prevalence of strangles in horses on the basis of PCR
was recorded and it was found to be the highest during the months of February,
March, April and May as shown in fig. 4.3. While few cases were seen during the
months of January, June and July and no cases were seen during the remaining
months. The differences among prevalence rates during different months of year were
found to be significant (p<0.05). It was concluded from the present study, that the
strangles season in Pakistan from January to July is at the peak. When we compared
strangles in different seasons (summer, winter, spring and autumn), the highest
prevalence was recorded during spring season. The reason may be due to maximum
exposure of horses during the spring time.
Age Groups
n Number positive for S. equi Total
(%) Jan Feb Mar Apr May Jun July Aug Sep Oct Nov Dec <1year 50 02 09 11 14 07 02 01 00 00 00 00 00 46(92)
1-2year 50 01 09 15 12 02 04 00 00 00 00 00 00 43(86)
2-3year 50 00 03 06 04 04 00 00 00 00 00 00 00 17(34)
3-4year 50 00 01 04 03 02 00 00 00 00 00 00 00 10(20)
4-5year 50 00 01 01 03 01 00 00 00 00 00 00 00 06(12)
Total 250 03 23 37 36 16 06 01 00 00 00 00 00 122(48.8)
53
Fig 4.3: Month wise prevalence of Strangles in horses on the basis of PCR.
Prevalence of Strangles in nasal discharge:
The prevalence of strangles in nasal discharge of horses on the basis of PCR
in different age groups and in different months of year is shown in table 4.5. The
prevalence rate in less than 1 year of age was 30(90.91%), 1-2 years of age was
31(83.78%), 2-3 years of age was 7(20%), 3-4 of years age 6(13.33%) and 4-5 years
of age was 3(6.82%). The highest prevalence in nasal discharge samples was recorded
in less than 1 year of age then 1-2 years and so on.
54
Table 4.5: Prevalence of Strangles in nasal discharge of horses on the basis of PCR.
Age
Groups n
Number positive for S. equi Total (%) Jan Feb Mar Apr May Jun July Aug Sep Oct Nov Dec
<1year 33 01 04 08 10 04 02 01 00 00 00 00 00 30(90.91)
1-2year 37 01 05 11 09 01 04 00 00 00 00 00 00 31(83.78)
2-3year 35 00 01 03 01 02 00 00 00 00 00 00 00 07(20.00)
3-4year 45 00 01 01 02 02 00 00 00 00 00 00 00 06(13.33)
4-5year 44 00 01 00 01 01 00 00 00 00 00 00 00 03(06.82)
Total 194 02 12 23 23 10 06 01 00 00 00 00 00 77(39.69)
Chi-square analysis showed significant difference in prevalence of strangles among all age groups (Chi-square value 104.81, P- value 0.0001)
Prevalence of Strangles in pus samples of sub-mandibular lymph nodes:
The prevalence of strangles in pus samples of the sub-mandibular lymph
nodes of horses on the basis of PCR in different age groups and in different months of
year is shown in table 4.6. The prevalence rate under 1 year of age 16(94.12%), 1-2
years of age 12(92.31%), 2-3 years of age 10(66.67%), 3-4 years of age 4(80%) and
4-5 years of age was 3(50%). The highest prevalence in pus samples of sub-
mandibular lymph nodes of horses was recorded in less than 1 year of age and then
between 1-2 years and so on.
When compared the prevalence of strangles in nasal discharge and pus
samples of sub-mandibular lymph nodes in different age groups. It was observed that
the number of cases of strangles were significantly higher (p<0.05) in pus samples
from sub-mandibular lymph nodes as compare to nasal discharge samples.
55
Table 4.6: Prevalence of Strangles in pus samples of sub-mandibular lymph nodes of horses on the basis of PCR.
Age
Groups n
Number positive for S. equi Total (%) Jan Feb Mar Apr May Jun July Aug Sep Oct Nov Dec
<1year 17 01 05 03 04 03 00 00 00 00 00 00 00 16(94.12)
1-2year 13 00 04 04 03 01 00 00 00 00 00 00 00 12(92.31)
2-3year 15 00 03 03 02 02 00 00 00 00 00 00 00 10(66.67)
3-4year 05 00 00 02 02 00 00 00 00 00 00 00 00 04(80.00)
4-5year 06 00 00 01 02 00 00 00 00 00 00 00 00 03(50.00)
Total 56 01 12 13 13 06 00 00 00 00 00 00 00 45(80.36)
Chi-square analysis showed non significant difference in prevalence of strangles among all age groups (Chi-square value 08.50, P- value 0.075)
Comparison of Culture and PCR:
From the results of present study it was observed that the sensitivity of the
Polymerase Chain Reaction appears to be much greater, than the culture for both
nasal and pus samples taken from the affected submandibular lymph nodes. Since the
test is completed in four to five hours it can be an effective tool in the control and
management of outbreaks. Finally it was concluded that culture along with PCR is the
best technique for the diagnosis of strangles, because the culture is of value as it
definitively establishes infection and can conveniently be performed on the same
samples used for PCR.
56
PREVALENCE OF STRANGLES IN MULES:
Prevalence of Strangles in mules on the basis of culture:
The prevalence of strangles in mules on the basis of culture in different age
groups and in different months of year is shown in table 4.7. The overall prevalence
rate in less than 1 year of age 41(82%), 1-2 years of age 39(78%), 2-3 years of age
10(20%), 3-4 years of age 6(12%) and 4-5 years of age was 3(6%). The highest
prevalence was recorded in less than 1 year of age then 1-2 years and decreased with
increase in age. The difference between prevalence rates among mules of different
age groups was significant (p<0.05). Therefore it is concluded from the result of
present study that the mules under 2 years of age were highly susceptible to strangles
infection as compared to over 2 years of age.
Table 4.7: Prevalence of Strangles in nasal discharge and pus samples of sub-mandibular lymph nodes of mules on the basis of culture.
Chi-square analysis showed significant difference in prevalence of strangles among all age groups (Chi-square value 115.96, P- value 0.0001)
Year round prevalence of strangles in mules was also recorded and it was
found to be the highest during the months of February, March, April and May.
Age Groups
n Number positive for S. equi Total
(%) Jan Feb Mar Apr May Jun July Aug Sep Oct Nov Dec <1year 50 00 07 11 17 06 00 00 00 00 00 00 00 41(82%)
1-2year 50 00 06 10 19 04 00 00 00 00 00 00 00 39(78%)
2-3year 50 00 01 02 05 02 00 00 00 00 00 00 00 10(20%)
3-4year 50 00 00 01 04 01 00 00 00 00 00 00 00 06(12%)
4-5year 50 00 00 00 03 00 00 00 00 00 00 00 00 03(06%)
Total 250 00 14 24 48 13 00 00 00 00 00 00 00 99(39.6)
57
Results are shown in fig. 4.4. While few cases were seen during the months of
January, June and July and no cases were seen during remainding months.
The significant difference was also observed (p<0.05) among prevalence rates
of different months of year. It was concluded that the strangles season in Pakistan
starts from month of January through the month of July, which was similar to the
horses, results. Similarly, when comparing the prevalence of strangles in different
seasons of Pakistan (summer, winter, spring and autumn), the highest prevalence rates
were recorded during the months of spring.
Fig 4.4: Month wise prevalence of Strangles in mules on the basis of culture.
Prevalence of Strangles in nasal discharge:
The prevalence of strangles in nasal discharge of mules on the basis of culture
in different age groups and in different months of year is shown in table 4.8. The
prevalence rate under 1 year of age 23(74.2%), 1-2 years of age 26(74.3%), 2-3 years
of age 4(10.30%), 3-4 years of age 4(9.80%) and 4-5 years of age was 2(4.80%). The
58
highest prevalence rate in nasal discharges of mules was recorded in 1-2 years of age
and then under 1 year of age while the lowest prevalence rate was recorded in 4-
5years of age that is also similar to horses.
Table 4.8: Prevalence of Strangles in nasal discharge of mules on the basis of culture.
Age
Groups n
Number positive for S. equi Total (%) Jan Feb Mar Apr May Jun July Aug Sep Oct Nov Dec
<1year 31 00 03 07 11 02 00 00 00 00 00 00 00 23(74.2)
1-2year 35 00 03 07 14 02 00 00 00 00 00 00 00 26(74.3)
2-3year 39 00 00 01 02 01 00 00 00 00 00 00 00 04(10.3)
3-4year 41 00 00 01 02 01 00 00 00 00 00 00 00 04(09.8)
4-5year 42 00 00 00 02 00 00 00 00 00 00 00 00 02(04.8)
Total 188 00 06 16 31 06 00 00 00 00 00 00 00 59(31.4)
Chi-square analysis showed significant difference in prevalence of strangles among all age groups (Chi-square value 87.11, P- value 0.0001)
Prevalence of Strangles in pus samples of sub-mandibular lymph nodes:
The prevalence of strangles in pus samples of sub-mandibular lymph nodes of
mules on the basis of culture in different age groups and in different months of year is
shown in table 4.9. The prevalence rate under 1 years of age 18(94.70%), 1-2 years of
age was 13(86.70%), 2-3 years of age was 6(54.5%), 3-4 years of age 2(22.20%) and
4-5 years of age was 1(12.50%) on the basis of culture. The highest prevalence in pus
samples of sub-mandibular lymph nodes of mules was recorded in the group having
less than 1 year of age and then between 1-2 years and so on.
The prevalence of strangles in nasal discharge and pus samples of sub-
mandibular lymph nodes was compared in different age groups. It was observed that
59
number of cases of strangles were significantly higher (p<0.05) in pus samples from
sub-mandibular lymph nodes as compared to nasal discharge samples.
Table 4.9: Prevalence of Strangles in pus samples of sub-mandibular lymph nodes of mules on the basis of culture.
Age Groups
n Number positive for S. equi Total
(%) Jan Feb Mar Apr May Jun July Aug Sep Oct Nov Dec <1year 19 00 04 04 06 04 00 00 00 00 00 00 00 18(94.7)
1-2year 15 00 03 03 05 02 00 00 00 00 00 00 00 13(86.7)
2-3year 11 00 01 01 03 01 00 00 00 00 00 00 00 06(54.5)
3-4year 09 00 00 00 02 00 00 00 00 00 00 00 00 02(22.2)
4-5year 08 00 00 00 01 00 00 00 00 00 00 00 00 01(12.5)
Total 62 00 08 08 17 07 00 00 00 00 00 00 00 40(64.5)
Chi-square analysis showed significant difference in prevalence of strangles among all age groups (Chi-square value 27.75, P- value 0.0001)
Prevalence of Strangles on the basis of Polymerase Chain Reaction:
The overall prevalence of strangles in mules on the basis of Polymerase Chain
Reaction in different age groups and in different months of year is shown in table
4.10. The prevalence rate under 1 year of age was 44(88%), 1-2 years of age
42(84%), 2-3 years of age 13(26%), 3-4 years of age 8(16%) and 4-5 years of age was
6(12%). The highest prevalence was recorded in less than 1 year of age and then
between 1-2 year and decreased with increase in age. A significant difference was
observed (p<0.05) between prevalence rates, among mules of different age groups.
60
Table 4.10: Prevalence of Strangles in nasal discharge and pus samples of sub-mandibular lymph nodes of mules on the basis of PCR.
Chi-square analysis showed significant difference in prevalence of strangles among all age groups (Chi-square value 114.26, P- value 0.0001) Prevalence of strangles in mules, during different months of the year, was also
recorded. It was found to be the highest during the months of February, March, April
and May and is shown in fig. 4.5. While few cases were seen during the months of
January, June and July and no cases were seen during rest of the months. The results
of different months were significantly different (p<0.05) from each other.
Fig 4.5: Month wise prevalence of Strangles in mules on the basis of PCR.
Age Groups
n Number positive for S. equi Total
(%) Jan Feb Mar Apr May Jun July Aug Sep Oct Nov Dec <1year 50 01 07 12 18 06 00 00 00 00 00 00 00 44(88%)
1-2year 50 00 08 11 19 04 00 00 00 00 00 00 00 42(84%)
2-3year 50 00 02 04 05 02 00 00 00 00 00 00 00 13(26%)
3-4year 50 00 01 02 04 01 00 00 00 00 00 00 00 08(16%)
4-5year 50 00 02 01 03 00 00 00 00 00 00 00 00 06(12%)
Total 250 01 20 30 49 13 00 00 00 00 00 00 00 113(45.2)
61
Prevalence of Strangles in nasal discharge:
The prevalence of strangles in nasal discharge of mules on the basis of PCR in
different age groups and in different months of year is shown in table 4.11. The
prevalence rate under 1 year of age was 26(83.9%), between 1-2 years of age was
28(80.0%), between 2-3 years of age was 7(17.90%), between 3-4 years of age
4(9.80%) and 4-5 years of age was 4(9.50%) on the basis of PCR. The highest
prevalence in nasal discharge of mules was recorded in the group having less than 1
year of age and then 1-2 years of age.
Table 4.11: Prevalence of Strangles in nasal discharge of mules on the basis of PCR.
Age
Groups n
Number positive for S. Equi Total (%) Jan Feb Mar Apr May Jun July Aug Sep Oct Nov Dec
<1year 31 01 03 08 12 02 00 00 00 00 00 00 00 26(83.9)
1-2year 35 00 05 07 14 02 00 00 00 00 00 00 00 28(80.0)
2-3year 39 00 01 02 02 02 00 00 00 00 00 00 00 07(17.9)
3-4year 41 00 00 01 02 01 00 00 00 00 00 00 00 04(09.8)
4-5year 42 00 01 01 02 00 00 00 00 00 00 00 00 04(09.5)
Total 188 01 10 19 32 07 00 00 00 00 00 00 00 69(36.7)
Chi-square analysis showed significant difference in prevalence of strangles among all age groups (Chi-square value 90.04, P- value 0.0001)
Prevalence of Strangles in pus samples of sub-mandibular lymph nodes:
The prevalence of strangles in pus samples of sub-mandibular lymph nodes of
mules on the basis of PCR in different age groups and in different months of year is
shown in table 4.12. The prevalence rate under 1 year of age was 18(94.70%),
between 1-2 years of age was 14(93.3%), between 2-3 years of age was 6(54.5%),
62
between 3-4 years of age 4(44.40%) and 4-5 years of age was 2(25.0%) on the basis
of PCR. The highest prevalence in pus samples of sub-mandibular lymph nodes of
mules was recorded in the group aging less than 1 year and then between 1-2 years
and so on.
The prevalence of strangles in nasal discharge and pus samples of sub-
mandibular lymph nodes was compared in different age groups. It was observed that
number of cases of strangles were significantly higher (p<0.05) in pus samples from
sub-mandibular lymph nodes as compared to nasal discharge samples.
Table 4.12: Prevalence of Strangles in pus samples of sub-mandibular lymph nodes of mules on the basis of PCR
. Age
Groups n
Number positive for S. Equi Total (%) Jan Feb Mar Apr May Jun July Aug Sep Oct Nov Dec
<1year 19 00 04 04 06 04 00 00 00 00 00 00 00 18(94.7)
1-2year 15 00 03 04 05 02 00 00 00 00 00 00 00 14(93.3)
2-3year 11 00 01 01 03 01 00 00 00 00 00 00 00 06(54.5)
3-4year 09 00 01 00 03 00 00 00 00 00 00 00 00 04(44.4)
4-5year 08 00 01 00 01 00 00 00 00 00 00 00 00 02(25.0)
Total 62 00 10 09 18 07 00 00 00 00 00 00 00 44(71.0)
Chi-square analysis showed significant difference in prevalence of strangles among all age groups (Chi-square value 21.57, P- value 0.0001)
Comparison of Culture and PCR in Mules:
The results of present study showed that the sensitivity of the Polymerase Chain
Reaction appears to be much greater than culture for both nasal and pus samples
taken from affected submandibular lymph nodes. The PCR can be completed in four
to five hours, might be an effective tool in the control and management of outbreaks.
63
Finally it was concluded that culture along with PCR is the best technique for the
diagnosis of strangles because culture is of value as it definitively establishes
infection and can conveniently be performed on the same samples used for PCR.
Fig.4.6: PCR amplification of DNAs from equine isolates of S. equi with specific primers. Lanes 1-9 shows the PCR products of S. equi with sized 677 and 325 bp by
using SeM primers. Lane 10 and 11 serve as -ve control and +ve control (CF32, S. equi, USA,) respectively.
64
MORTALITY:
Mortality rate in horses:
Mortality rate in horses suffering from strangles is shown in table 4.13, out of
122 horses only 2(1.64%) horses died, one from less than one year of age and one
from 1-2 years of age. The difference in mortality rate of affected horses among
different age groups was not significant (P>0.05) however it is concluded from the
present study that the severity of strangles is greater in horses of less than 2 years of
age, as compared to over two year of age.
Table 4.13: Mortality rate in horses under 5 years of age
Age groups Number affected
Mortality Number %age
<1year 46 01 2.17
1-2year 43 01 2.32
2-3year 17 00 00.0
3-4year 10 00 00.0
4-5year 06 00 00.0
Total 122 02 1.64
65
Mortality rate in Mules:
Similarly mortality rate in mules with strangles is shown in table 4.14, out of
113 mules, only 1(0.88%) mule that was from less than one year of age died. The non
significant difference (P>0.05) was observed in mortality rate of affected mules of
different age groups. It is concluded from the results of the present study that severity
of strangles was greater in the mules of less than 2 year of age as compared to over
two years of age.
Table 4.14: Mortality rate in mules under 5 years of age
Age groups Number affected
Mortality Number %age
<1year 44 01 2.27
1-2year 42 00 00.0
2-3year 13 00 00.0
3-4year 08 00 00.0
4-5year 06 00 00.0
total 113 01 0.88
When comparing the mortality rate in horses and mules no significant
difference (P>0.05) was observed among all age groups however it was known from
the present study that the severity of diseases was seen more in horses as compared to
mules.
66
Variations in SeM, SzPSe and Se 18.9 proteins of S.equi:
SeM.
Results of PCR of the SeM genes of 24 selected isolates of S. equi and the US
prototype strain CF32 generated products of 1812 bp is shown in fig. 4.7. Resulting
DNA sequences encoded 19 different SeM alleles including numbers 71-76 not
previously included in the database (www.pubmlst.org/szooepidemicus/) (Table
4.16). Residues most frequently subjected to substitution were located at position 58,
63, 108 and 143. Single nucleotide polymorphisms (SNPs) in SeM were found at 93
loci and totalled 181. Fifty-eight of these were non-synonymous, that is, mutations
were resulting in amino acid replacements in SeM. Non-synonymous SNPs were 15.7
times more frequent in the N-terminal region (positions 114 to 629) than in the
remainder of the SeM sequence (Table 4.18). Phylogenies generated by
neighbourhood joining indicate that SeM allele 71 identified in isolates S24 and L32
from Pakistan was the most distantly related allele of the 25 isolates in the study. This
is explained by the remote and isolated Pakistani equid population, which
hypothetically favors preservation and continued divergence of a specific SeM allele.
Other newly identified alleles (72 - 76) in N. America isolates showed 96 - 99%
similarity with alleles 62, 59, 57 and 37 in the SeM database.
67
Table 4.15: Details of all amplicons used in the present study.
Strain ID Year isolated Geographic OriginAmplicon detected
SeM SzPSe Se18.9 EqbES24 2010 Pakistan + + + +
L32 2009 Pakistan + + + +
MN08 1998 USA(MN) + + + +
119 1191 Scotland + + + +
JBa 1998 USA (NY) + + + +
JBb 1996 USA (KY) + + + +
FF 1998 USA (KY) + + + +
836a 1997 USA (KY) + + + +
836b 1998 USA (KY) + + + +
LEX 1990 USA (KY) + + + +
E33 1976 USA (NY) + + + +
181 1990 AUSTRALIA + + + +
331 1982 Swedish + + + +
BM 1983 Ireland + + + +
M4 1991 Germany + + + +
E9 1976 USA (NY) + + + +
FT 2003 USA (IA) + + + +
JP 2010 Denmark + + + +
NM 2007 USA (KY) + + + +
MER7 2007 USA (VA) + + + +
15C 2001 USA (UT) + + + +
Lou 2010 USA (ME) + + + +
107 1991 Scotland + + + +
K58 1969 USA (KY) + + + +
W60 1976 USA (NY) + + - -
CF32 1981 USA (NY) + + + +
68
Table 4.16: SeM alleles in S. equi isolated over a period of 40 years in N. America, Europe and Asia.
Isolate Allele No.
Amino acid residues s 38 47 51 52 53 56 57 58 60 62 63 65 69 90 99 102 103 104 106 107 108 110 111 113 122 125 127 143
TW 1 S R D L K L S E A S R A Q L R Y Y N L M H S S L R S A S CF32 2 * * * * * * * D * * G * * * * * * * * * * * * * * * * R 4047 3 * * * * * * * * * * * * * * * * * * * V * * * * * * * * S24 71 * T * F * * N * * T G * * * * * * * * I Q * * P S N S * L32 71 * T * F * * N * * T G * * * * * * * * I Q * * P S N S *
MN08 2 * * * * * * * D * * G * * * * * * * * * * * * * * * * * 119 1 * * * * * * * * * * * * * * * * * * * * * * * * * * * * JBa 26 * * * * * * * D * * * * * * * * * * * * R * L * * * * R JBb 42 * K N * R F * * * * * * * * * * N * * * R P P * S * * * FF 27 * K N * R F * * * * * * * * * * N * * * R P P * S * * R
836a 31 * * * * * * * D * * G * * * * F * * * * * * * * * * * R 836b 32 * * * * * * * D * * G * * * * F H * * * R * * * * * * R
Lex90 2 * * * * * * * D * * G * * * * * * * * * * * * * * * * R E33 22 P * * * * * * * * * G * * * * * * * * K * P * * * * * R 181 15 * * * * * * * * * * * * * * * * * * * * Q * * * * * * * 331 36 * * * * * * * D * * * * L * * * * * * * * * * * * * * * BM 23 P * * * * * * * * * G * * * * * H * * * * * * * * * * R M4 72 * * * * * * * D * G * * * * * * * * * * * * * * * * * * E9 22 P * * * * * * * * * G * * * * * * * * K * P * * * * * R FT 28 * * * * * * * D * * * * * * * * * * * * * * * * * * * R JP 73 * * * * * * * * D * * T * F * * * * * * R * * * * * * *
NM 39 * * * * * * N D * * * * * * K * * * * * * * * * * * * * MER7 74 * * * * * * * * * * * * * * * * H Y S * * P * S * * * R 15C 75 * * * * * * N D * * * * * * K * * * * * R * * * * * * * Lou 2 * * * * * * * D * * G * * * * * * * * * * * * * * * * R 107 76 * * * * * * * * * * * * * * * * * * * R A F * * * * * * K58 2 * * * * * * * D * * G * * * * * * * * * * * * * * * * R
69
Fig.4.7:. PCR amplification of DNAs from equine isolates of S. equi to analyse variation in SeM (a), SzPSe (b), Se18.7 (c) and EqbE (d). Lanes 1 -11 included PAK24 (Pakistan), 1079 (Scotland), K-58 (USA), Boldmani (Ireland), 181093SYD (Australia), German Martin-4 (Germany), John Paul (Denmark), Lexington 90 (Lexington, KY, USA), 331(81) (Sweden), S. zooepidemicus W60 (-ve control) and S. equi CF32 (+ve control).
70
Table 4.17: Identification of six new alleles by BLAST analysis against www.pubmlst.org/szooepidemicus/
Strain Peptide having allelic
residues
Homology Result
Pak24
SEVSRTATPTLSRDFKNRLNEIAITGDHAS
SAQKVRNLLKGASVRDLQALLRGLDSARAA
YGRDDYYNLLIQLSSMPNDKPDGDRSQLNL
SSLLVDEIEKRIADGDSYAK
allele 41 - 90.00%
allele 40 - 90.00%
allele 35 - 88.18%
New allele 1
Pak32
SEVSRTATPTLSRDFKNRLNEIAITGDHAS
SAQKVRNLLKGASVRDLQALLRGLDSARAA
YGRDDYYNLLIQLSSMPNDKPDGDRSQLNL
SSLLVDEIEKRIADGDSYAK
allele 41 - 90.00%
allele 40 - 90.00%
allele 35 - 88.18%
New allele 1
German
Martin4
SEVSRTATPRLSRDLKNRLSDIAIGRDAS
SAQKVRNLLKGASVGDLQALLRGLDSARA
AYGRDDYYNLLMHLSSMLNDKPDGDRRQL
SLASLLVDEIEKRIADGDS
allele 62 - 99.06%
allele 51 - 98.11%
allele 69 - 98.11%
New allele 2
John Paul
SEVSRTATPRLSRDLKNRLSEIDISRDTS
SAQKVRNLLKGASVGDLQALLRGFDSARA
AYGRDDYYNLLMRLSSMLNDKPDGDRRQL
SLASLLVDEIEKRIADGDS
allele 59 - 99.06%
allele 6 - 98.11%
allele 47 - 97.17%
New allele 3
Mergana
Aug 07
SEVSRTATPRLSRDLKNRLSEIAISRDAS
SAQKVRNLLKGASVGDLQALLRGLDSARA
AYGRDDYHNLSMHLPSMSNDKPDGDRRQL
SLASLLVDEIEKRIADGDR
allele 57 - 96.23%
allele 25 - 96.23%
allele 54 - 96.19%
New allele 4
15C
SEVSRTATPRLSRDLKNRLNDIAISRDAS
SAQKVRNLLKGASVGDLQALLRGLDSARA
AYGKDDYYNLLMRLSSMLNDKPDGDRRQL
SLASLLVDEIEKRIADGDS
allele 39 - 99.06%
allele 38 - 98.11%
allele 61 - 97.17%
New allele 5
1079
SEVSRTATPRLSRDLKNRLSEIAISRDASS
AQKVRNLLKGASVGDLQALLRGLDSARAAY
GRDDYYXFLRAFFSMLKDKPAGDFRQLSLA
‘SLLVDEIEKRIADGDS
allele 15 - 91.51%
allele 43 - 91.51%
allele 1 - 91.51%
New allele 6
71
Table 4.18: Frequency of single nucleotide polymorphism (SNPs) in SeM, SzPSe and Se18.9 in 25 isolates of S. equi.
SzPSe.
PCR of the SzPSe genes of 24 selected isolates of S. equi and the US
prototype strain CF32 generated products of 1152 bp is shown in fig. 4.7. With one
exception, sequence analysis of the SzPSe genes provided no instances of variation.
The single exception, Australian isolate 181, had a deletion of one PEPK repeat.
Remarkably, although 92 SNPs were found at 48 loci in SzPSe of the 25 S. equi
isolates including CF32 S. equi, no SNPs encoding non-synonymous substitutions
were found (Table 4.18). Thus, there is evidence of a high rate of recombination in
the SzP gene.
Moreover, the HV region appears to have been horizontally acquired since its
G-C % (38.3) differs significantly (p<0.01) from that (47.0) of the remaining SzPSe
sequence. Recombination and the presence of exogenous DNA sequence are factors
that would favour occurrence of SNPs. The biological/immunological significance of
this variation is not understood, but does not appear to involve opsonogenic epitopes.
Future work might logically address the effect of variation on the conformational
adhesion epitope on host cell specificity.
Se 18.9.
PCR of the Se18.9 genes of 24 selected isolates of S. equi and the US
prototype strain CF32 generated products of 481 bp is shown in fig. 4.7. Se 18.9
Gene Bases SNP Loci SNPs Non-synonymous
SNPs (%)SeM Nucleotide 114 - 429 315 44 97 46 (47) Nucleotide 430 - 1605 1290 49 84 12 (14) SzPSe 1140 48 92 0 (0) Se18.9 492 02 04 0 (0)
72
protein has a proven virulence function in common with SeM, it is logically a target
of immune selection pressure yet only 2 SNP loci were found in the DNA sequences
of the Se18.9 genes in 25 isolates. The unexpected absence of variants of Se18.9 in a
population of SeM allelic variants of S. equi argues either for an immutable and
essential structure or virulence function that is minor compared to that of SeM. It
might also be argued that as a secreted protein, Se18.9 might have less survival value
for S. equi than a protein anchored on its surface. The much lower frequency of SNP
loci in Se18.9 compared to SeM and SzPSe is unexplained.
Table 4.19: Details of SzPSe of all isolates of S. equi S. equi strain ID N-terminus HV region PE(P)K repeats
S24 N2 HV4 18 L32 N2 HV4 18
MN08 N2 HV4 18 119 N2 HV4 18 JBa N2 HV4 18 JBb N2 HV4 18 FF N2 HV4 18
836a N2 HV4 18 836b N2 HV4 18 LEX N2 HV4 18 181 N2 HV4 17 E33 N2 HV4 18 331 N2 HV4 18 BM N2 HV4 18 M4 N2 HV4 18 E9 N2 HV4 18 FT N2 HV4 18 JP N2 HV4 18
NM N2 HV4 18 MER7 N2 HV4 18 15C N2 HV4 18 Lou N2 HV4 18 107 N2 HV4 18 K58 N2 HV4 18 W60 N2 HV4 18 CF32 N2 HV4 18
73
STUDY OF CARRIER STATUS OF HORSES AND MULES:
Carrier status of Horses:
Out of 122 horses, 20 horses (10<2 years and 10 between 2 and 5 years of
age) positive for strangles agent after one week of infection and were monitored for
12 weeks to study their carrier status is shown in table 4.20. After the end of the 3rd
week all horses < 2 years of age were positive but at the end of 4th to 7th weeks there
remained 5, 2, 1 and zero out of 10, respectively on the basis of culture, whereas
through polymerase chain reaction at the end of the 4th week all horses <2 years of
age were positive, but at the end of 5th to 10th weeks there remained 7, 5, 4, 2, 1 and
zero out of 10, respectively. While in 2 and 5 years old horses, all were positive up to
the 1st week but at the end of 2nd to 8th weeks there were 9, 7, 6, 3, 1, 1 and zero
horses positive, respectively out of 10, on the basis of culture. Through PCR all
horses were positive up to 4th week but at the end of 5th to 9th weeks there were 9, 7,
6, 3, 2 and zero. Horses were declared free of infection on the basis of three
consecutive negative samples through culture and PCR.
Therefore it is concluded from the findings of present study that sensitivity of
Polymerase Chain Reaction appears to be much greater than culture for detection of
carrier status of horses. According to present study, recovered horses should be kept
in quarantine period at least for 9 weeks because the recovered horses remain in
shedder state for a prolonged period of time, and through periodic shedding of S equi
can be a source of infection for susceptible equines.
74
Table 4.20: Comparison of culture and PCR for identification of carrier of S. equi in naturally infected horses ≤ 5 years of age.
Weeks post infection n=20 Number positive for S.equi
<2year 2-
5year <2year 2-5year
Culture PCR Culture PCR Zero/Infection day 10 10 10 10 10 10
1st week post
infection 10 10 10 10 10 10
2nd week post
infection 10 10 10 10 09 10
3rd week post
infection 10 10 10 10 07 10
4th week post
infection 10 10 05 10 06 10
5th week post
infection 10 10 02 07 03 09
6th week post
infection 10 10 01 05 01 07
7th week post
infection 10 10 - 04 01 06
8th week post
infection 10 10 - 02 - 03
9th week post
infection 10 10 - 01 - 02
10th week post
infection 10 10 - - - -
11th week post
infection 10 10 - - - -
12th week post
infection 10 10 - - - -
75
Carrier status of Mules:
Similarly, out of 113 mules, 20 mules (10<2 years and 10 between 2 and 5
years of age) remaining positive after one week of infection were monitored for 12
weeks to study their carrier status. Results are shown in table 4.21. After the end of
2nd week all mules < 2 years of age were positive but at the end of 3rd to 6th weeks
there remained 7, 3, 1 and zero mules out of 10, respectively on the basis of culture.
Through the polymerase chain reaction at the end of the 5th week all mules <2 years
of age were positive, but at the end of 6th to 10th weeks there remained 9, 7, 3, 2 and
zero mules out of 10, respectively.
While in the group having 2 to 5 year old mules, all were positive up to the 2nd
week but at the end of 3rd to 7th weeks there were 6, 4, 2, 1, 1 and zero mules out of
10 mules, respectively on the basis of culture. Through PCR, all mules were positive
up to 5th week but at the end of 6th to 10th weeks there were 8, 5, 2, 1 and zero
respectively. Mules were declared free of infection on the basis of three consecutive
negative samples through culture and PCR.
From the results of the present study, it can therefore be concluded that
sensitivity of Polymerase Chain Reaction is much greater than culture for the study of
carrier status of mules. Moreover, recovered mules should be kept in the quarantine at
least for a period of 9 weeks, because the recovered mules remains carrier for
prolonged period of time and through periodic shedding of S equi can cause infection
in apparently healthy but susceptible animals.
76
Table 4.21: Comparison of culture and PCR for identification of carrier of S. equi in naturally infected mules ≤ 5 years of age.
Weeks post infection n=20 Number positive for S.equi
<2year 2-
5year <2year 2-5year
Culture PCR Culture PCR Zero/Infection day 10 10 10 10 10 10
1st week post
infection 10 10 10 10 10 10
2nd week post
infection 10 10 10 10 10 10
3rd week post
infection 10 10 07 10 06 10
4th week post
infection 10 10 03 10 04 10
5th week post
infection 10 10 01 10 02 10
6th week post
infection 10 10 - 09 01 08
7th week post
infection 10 10 - 07 01 05
8th week post
infection 10 10 - 03 - 02
9th week post
infection 10 10 - 02 - 01
10th week post
infection 10 10 - - - -
11th week post
infection 10 10 - - - -
12th week post
infection 10 10 - - - -
77
PHASE 4:
HAEMATOLOGICAL EXAMINATION:
Haematological studies were conducted to examine the effect of strangles on
various blood parameters during active infection and carrier state in horses and mules.
1-Haematological Examination of Diseased horses and mules:
For haematological examination 40 horses (n=20 healthy horses; n= 20
diseased horses) and 40 mules (n=20 healthy mules; n= 20 diseased mules) under 5
years of age suffering from natural outbreak of strangles from Lahore and Sargodha
districts of Punjab province of Pakistan were included. Parameters like total white
blood cell count, mean segmented neutrophils count, total lymphocytic count, total
monocytic count, total eosinophilic count, total basophilic count, total erythrocytic
count, packed cell volume and haemoglobin concentration were studied.
Total white blood cell count:
Total white blood cell count in horses and mules suffering for strangles is
shown in table 4.22. In horses total WBCs were 14.99±0.22x109/L at 1st week,
14.07±0.19x109/L at 2nd week, 12.80±0.16x109/L at 3rd week and 11.91±0.16x109/L
at 4th week post infection. When compared total WBCs of diseased and healthy
horses on weekly basis significant increase was observed up to 3rd week of infection
while during the 4th week of infection there was no significant increase (P>0.05).
Among all four weeks total WBCs were according to the following order 4th
<3rd<2nd<1st week. It was also observed that at the end of 1st week post infection
total white blood cell count was highest which significantly decreased to the normal
values during the subsequent weeks (P<0.05).
78
In mules total white blood cell count was 13.27±0.22x109/L at the end of 1st
week, 12.98±0.22x109/L at 2nd week, 12.18±0.17x109/L at 3rd week and
10.89±0.19x109/L at 4th week post infection. Similar pattern was observed in mules
as in horses when compared diseased mules with healthy mules on weekly basis
significant increase was observed up to 3rd week of infection while during the 4th
week of infection there was no significant increase (P>0.05). It was also observed that
rise in total white blood cell count was rapid and much higher in horses as compared
to mules.
Table 4.22: Total white blood cell count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
Mean in a column followed by the same letter were not significantly different at P≤0.05, by Tukey HSD test. * indicates significant difference (p<0.05) among healthy and diseased groups ** indicates non-significant difference (p>0.05) between healthy and diseased group
When diseased horses compared with diseased mules, little difference was
observed, while increase in total white blood cell count was significantly higher
(P<0.05) in affected horses and mules than healthy animals as shown in fig. 4.8. It
was also observed that the level of total WBCs remained increased in affected
Post Infection Weeks
n Horses Mules
Horses Mules Healthy
n=20 Diseased
n=20 Healthy
n=20 Diseased
n=20
1st 40 40 *11.78±0.17a *14.99±0.22a *09.85±0.16b *13.27±0.22a
2nd 40 40 *10.71±0.16b *14.07±0.19b *10.73±0.09a *12.98±0.22a
3rd 40 40 *09.93±0.09c *12.80±0.16c *09.28±0.13c *12.18±0.17b
4th 40 40 **11.16±0.19b **11.91±0.16d **10.13±0.12b **10.89±0.19c
79
animals than healthy animals during all the four weeks. This indicates the
significance of total WBCs during disease condition and is one of the diagnostic
points of strangles disease.
Fig 4.8: Week wise comparison of Total white blood cell count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
Mean Segmented Neutrophilic Count (MSNC):
Data regarding mean segmented neutrophilic count in horses and mules
suffered from strangles is shown in table 4.23. In horses mean segmented neutrophilic
count was 07.07±0.11x109/L at 1st week, 06.98±0.07x109/L at 2nd week,
05.08±0.08x109/L at 3rd week and 04.55±0.07x109/L at 4th week post infection.
When compared diseased and healthy horses on weekly basis significant increase
(P<0.05) in the values was observed in all four weeks of infection. It was also seen
that at the end of 1st week post infection mean segmented neutrophilic count was
80
highest which significantly decreased to the normal values during the subsequent
weeks (P<0.05).
In mules mean segmented neutrophilic count was 06.80±0.10x109/L at 1st
week, 06.03±0.05x109/L at 2nd week, 05.45±0.07x109/L at 3rd week and
04.98±0.06x109/L at 4th week post infection. When the result of diseased mules
compared with healthy mules on weekly basis significant increase in the values was
observed up to 3rd week of infection while on the 4th week of infection the increase
was non significant (P>0.05).
When compared diseased horses with diseased mules, there were no
significant differences observed (P>0.05), while mean segmented neutrophilic count
significantly increased (P<0.05) in affected horses and mules as compared to healthy
animals as shown in fig. 4.9. It was also observed that the level of MSNC remained
increased in diseased animals than healthy animals during all the four weeks that
indicate significance of MSNC during disease condition and can also form one of the
diagnostic points of strangles disease.
Table 4.23: Mean segmented Neutrophilic count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
Mean in a column followed by the same letter were not significantly different at P≤0.05, by Tukey HSD test. * indicates significant difference (p<0.05) among healthy and diseased groups ** indicates non-significant difference (p>0.05) between healthy and diseased group
Post Infection Weeks
n Horses Mules
Horses Mules Healthy
n=20 Diseased
n=20 Healthy
n=20 Diseased
n=20
1st 40 40 *04.54±0.07a *07.07±0.11a *04.68±0.10a *06.80±0.10a
2nd 40 40 *04.72±0.08a *06.98±0.07a *03.98±0.04b *06.03±0.05b
3rd 40 40 *03.96±0.05b *05.08±0.08b *04.11±0.04b *05.45±0.07c
4th 40 40 *03.78±0.07b *04.55±0.07c **04.60±0.07a **04.98±0.06d
81
Fig 4.9: Week wise comparison of Mean segmented Neutrophilic count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
Total Lymphocytic count:
The result of total lymphocytic count in horses and mules suffering from
strangles is shown in table 4.24. Total lymphocytic count in horses were recorded as
02.71±0.05x109/L at the end of 1st week, 03.98±0.07x109/L at the end of 2nd week,
04.76±0.08x109/L at the end of 3rd week and 02.56±0.04x109/L at the end of 4th
week post infection. When diseased horses compared with healthy horses on weekly
basis there, no significant increase or decrease (P>0.05) during all four weeks was
observed.
In mules the values of total lymphocytic count was 03.89±0.05x109/L at 1st
week, 02.97±0.05x109/L at 2nd week, 03.03±0.04x109/L at 3rd week and
03.71±0.06x109/L at 4th week post infection. When compared diseased mules with
healthy mules on weekly basis a significant decrease (P<0.05) was observed in all
four weeks of infection and this was opposite to horses. It was further noticed that at
82
the end of 2nd week post infection total lymphocytic count was lowest which
significantly increased to the normal values during the subsequent weeks (P<0.05).
When data of diseased horses compared with that of diseased mules on weekly basis
there was no significant increase or decrease (P>0.05) observed among all four weeks
as shown in fig. 4.10.
Table 4.24: Total Lymphocytic count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
Mean in a column followed by the same letter were not significantly different at P≤0.05, by Tukey HSD test. * indicates significant difference (p<0.05) among healthy and diseased groups ** indicates non-significant difference (p>0.05) between healthy and diseased group
Fig 4.10: Week wise comparison of Total Lymphocytic count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
Post Infection Weeks
n Horses Mules
Horses Mules Healthy
n=20 Diseased
n=20 Healthy
n=20 Diseased
n=20 1st 40 40 **02.89±0.05c **02.71±0.05c *05.53±0.05a *03.89±0.05a
2nd 40 40 **03.48±0.06b **03.98±0.07b *05.40±0.03a *02.97±0.05b
3rd 40 40 **03.98±0.04a **04.76±0.08a *04.98±0.06b *03.03±0.04b
4th 40 40 **02.93±0.05c **02.56±0.04c *05.02±0.07b *03.71±0.06a
83
Total Monocytic count (TMC):
Data on total monocytic count in horses and mules suffered from strangles is
shown in table 4.25. Total monocytic count in horses was 00.53±0.021 x109/L at the
end of 1st week, 00.45±0.014 x109/L at the end of 2nd week, 00.49±0.011 x109/L at
the end of 3rd week and 00.47±0.011 x109/L at the end of 4th week. Comparison of
diseased horses with healthy horses revealed a non significant increase (P>0.05)
among all four weeks.
In mules the result of total monocytic count was 00.80±0.021x109/L at 1st
week, 00.87±0.014x109/L at 2nd week, 00.53±0.017 x109/L at 3rd week and
00.43±0.012 x109/L at 4th week post infection. When compared diseased mules with
healthy mules on weekly basis a significant increase (P<0.05) was observed in all
four weeks of infection, while in horses this increase was non significant (P>0.05).
When diseased horses compared with diseased mules on weekly basis the difference
of total monocytic count was observed non significant (P>0.05) among all four weeks
is shown in fig. 4.11. It was also observed that the value of total monocytic count
remained increased among all four weeks in diseased animals than healthy animals.
Table 4.25: Total Monocytic count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
Mean in a column followed by the same letter were not significantly different at P≤0.05, by Tukey HSD test. * indicates significant difference (p<0.05) among healthy and diseased groups ** indicates non-significant difference (p>0.05) between healthy and diseased group
Post Infection Weeks
N Horses Mules
Horses Mules Healthy
n=20 Diseased
n=20 Healthy
n=20 Diseased
n=20
1st 40 40 **00.36±0.018a **00.53±0.021a *00.17±0.010ab *00.80±0.021b
2nd 40 40 **00.38±0.020a **00.45±0.014b *00.20±0.009a *00.87±0.014a
3rd 40 40 **00.38±0.016a **00.49±0.011ab *00.18±0.010ab *00.53±0.017c
4th 40 40 **00.36±0.015a **00.47±0.011b *00.15±0.007b *00.43±0.012d
84
Fig 4.11: Week wise comparison of Total Monocytic count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
Total Eosinophilic count:
The results of total eosinophilic count in horses and mules suffering from
strangles is shown in table 4.26. Total eosinophilic count in horses was
00.48±0.010x109/L at 1st week, 00.32±0.022x109/L at 2nd week, 00.45±0.011 x109/L
at 3rd week and 00.50±0.014 x109/L at 4th week post infection. On comparison of
diseased horses with healthy horses on weekly basis a non significant difference
(P>0.05) of total eosinophilic count was observed.
Table 4.26: Total Eosinophil count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
Mean in a column followed by the same letter were not significantly different at P≤0.05, by Tukey HSD test. * indicates significant difference (p<0.05) among healthy and diseased groups ** indicates non-significant difference (p>0.05) between healthy and diseased group
Post Infection Weeks
N Horses Mules
Horses Mules Healthy
n=20 Diseased
n=20 Healthy
n=20 Diseased
n=20 1st 40 40 **00.53±0.016a **00.48±0.010a **00.27±0.014b **00.51±0.015a
2nd 40 40 **00.46±0.010c **00.32±0.022b **00.34±0.014a **00.54±0.015a
3rd 40 40 **00.51±0.014ab **00.45±0.011a **00.29±0.007b **00.49±0.007a
4th 40 40 **00.48±0.007bc **00.50±0.014a **00.28±0.007b **00.42±0.011b
85
Total eosinophilic count in mules was 00.51±0.015x109/L at 1st week,
00.54±0.015x109/L at 2nd week, 00.49±0.007 x109/L at 3rd week and 00.42±0.011
x109/L at 4th week. Similarly as in horses, the difference of total eosinophilic count in
diseased and healthy mules was observed non significant (P>0.05) among all four
weeks. When compared diseased horses with diseased mules on weekly basis the
difference of total eosinophilic count was also found non significant (P>0.05) as
shown in fig. 4.12.
Fig 4.12: Week wise comparison of Total Eosinophil count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
Total Basophil count:
Data regarding total basophilic count in horses and mules suffered from
strangles is shown in table 4.27. The results of total basophilic count in horses were
00.43±0.012x109/L at 1st week, 00.48±0.010x109/L at 2nd week, 00.20±0.007x109/L
at 3rd week and 00.07±0.003x109/L at 4th week of post infection. When compared
diseased and healthy horses on weekly basis significant increase was observed up to
86
3rd week of infection while on the 4th week of infection there was non significant
increase (P>0.05).
Table 4.27: Total Basophilic count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
Mean in a column followed by the same letter were not significantly different at P≤0.05, by Tukey HSD test. * indicates significant difference (p<0.05) among healthy and diseased groups ** indicates non-significant difference (p>0.05) between healthy and diseased group It was also observed that at the end of 2nd week post infection total basophilic
count was highest which significantly decreased to the normal values during the
subsequent weeks (P<0.05). Total basophilic count in mules were 00.17±0.010x109/L
at the end of 1st week, 00.13±0.007x109/L at the end of 2nd week, 00.20±0.007
x109/L at the end of 3rd week and 00.11±0.012x109/L at the end of 4th week post
infection. When diseased mules compared with healthy mules on weekly basis the
difference of total basophilic count was non significant (P>0.05). When the results of
diseased horses compared with diseased mules on weekly basis no significant
increase (P>0.05) was observed as shown in fig. 4.13.
Post Infection Weeks
N Horses Mules
Horses Mules Healthy
n=20 Diseased
n=20 Healthy
n=20 Diseased
n=20
1st 40 40 *00.05±0.004a *00.43±0.012b **00.05±0.003b **00.17±0.010b
2nd 40 40 *00.03±0.003b *00.48±0.010a **00.04±0.003b **00.13±0.007c
3rd 40 40 *00.04±0.003ab *00.20±0.007c **00.07±0.003a **00.20±0.007a
4th 40 40 **00.01±0.001c **00.07±0.003d **00.02±0.004c **00.11±0.012c
87
Fig 4.13: Week wise comparison of Total Basophilic count (x109/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
Total Erythrocytic count:
Data on total erythrocytes count in horses and mules suffering for strangles is
shown in table 4.28. Total erythrocyte count in horses was 5.97±0.04 x 1012/L at 1st
week, 6.03±0.04 x1012/L at 2nd week, 5.91±0.05 x 1012/L at 3rd week and 5.87±0.04
x 1012/L at 4th week. When compared diseased horses with healthy horses on weekly
basis the decrease of total erythrocytes count was observed non significant (P>0.05)
among all four weeks.
Total erythrocytic count in mules was 6.07±0.05 x 1012/L at the end of 1st
week, 6.35±0.05 x 1012/L at the end of 2nd week, 6.56±0.06 x 1012/L at the end of 3rd
week and 6.37±0.05 x 1012/L at the end of 4th week post infection. When compared
diseased mules with healthy mules on weekly basis the decreases of total erythrocytes
count was observed to be non significant (P>0.05) among all four weeks. Similarly
when diseased horses compared with diseased mules on weekly basis difference of
88
total erythrocytes count was observed non significant (P>0.05) among all four weeks
as shown in fig. 4.14.
Table 4.28: Erythrocytes count (X 1012/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
Mean in a column followed by the same letter were not significantly different at P≤0.05, by Tukey
HSD test.
* indicates significant difference (p<0.05) among healthy and diseased groups
** indicates non-significant difference (p>0.05) between healthy and diseased group
Fig 4.14: Week wise comparison of Erythrocytes count (X 1012/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
Post Infection Weeks
N Horses Mules
Horses Mules Healthy
n=20 Diseased
n=20 Healthy
n=20 Diseased
n=20
1st 40 40 **6.27±0.03a **5.97±0.04ab **6.79±0.07a **6.07±0.05c
2nd 40 40 **6.13±0.05ab **6.03±0.04a **6.81±0.07a **6.35±0.05b
3rd 40 40 **6.07±0.06b **5.91±0.05ab **6.77±0.06a **6.56±0.06a
4th 40 40 **6.02±0.04b **5.87±0.04b **6.80±0.07a **6.37±0.05ab
89
Packed cell volume:
Data regarding packed cell volume in horses and mules suffered from
strangles is shown in table 4.29. In horses packed cell volume was 29.14±0.14% at
the 1st week, 32.98±0.06% at the 2nd week, 30.67±0.17b % at the 3rd week
and28.10±0.09% at the end of 4th week post infection. When compared diseased and
healthy horses on weekly basis the decrease of packed cell volume was observed non
significant (P>0.05) among all four weeks.
In mules packed cell volume was 28.45±0.14% at the 1st week, 33.04±0.10%
at the 2nd week, 29.70±0.14% at the 3rd week and 31.13±0.23% at the end of 4th
week. When compared the data of diseased and healthy mules on weekly basis a non
significant (P>0.05) decrease of packed cell volume was observed among all four
weeks. Similarly the difference of values of packed cell volume in diseased horses
and diseased mules on weekly basis was also found to be non significant (P>0.05)
among all four weeks as shown in fig. 4.15.
Table 4.29: Packed cell volume (%) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
Mean in a column followed by the same letter were not significantly different at P≤0.05, by Tukey HSD test. * indicates significant difference (p<0.05) among healthy and diseased groups ** indicates non-significant difference (p>0.05) between healthy and diseased group
Post Infection Weeks
N Horses Mules
Horses Mules Healthy
n=20 Diseased
n=20 Healthy
n=20 Diseased
n=20 1st 40 40 **35.67±0.24b **29.14±0.14c **37.56±0.14a **28.45±0.14d
2nd 40 40 **34.89±0.17c **32.98±0.06a **36.70±0.07b **33.04±0.10a
3rd 40 40 **34.88±0.17c **30.67±0.17b **35.97±0.24c **29.70±0.14c
4th 40 40 **36.45±0.07a **28.10±0.09d **36.01±0.20c **31.13±0.23b
90
Fig 4.15: Week wise comparison of packed cell volume (%) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
Haemoglobin concentration:
The results of haemoglobin concentration in horses and mules suffering for
strangles are shown in table 4.30. Haemoglobin concentration in horses was
109.45±0.21g/L at 1st week, 100.67±0.11g/L at 2nd week, 99.12±0.20g/L at 3rd
week and 104.61±0.17g/L at 4th week post infection. When diseased and healthy
horses were compared on weekly, the decrease of haemoglobin concentration was
observed to be non significant (P>0.05) among all four weeks. It was also noted that
this decrease of haemoglobin concentration was more in mules than in horses.
91
Table 4.30: Haemoglobin concentration (g/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
Mean in a column followed by the same letter were not significantly different at P≤0.05, by Tukey HSD test. * indicates significant difference (p<0.05) among healthy and diseased groups ** indicates non-significant difference (p>0.05) between healthy and diseased group
Similarly haemoglobin concentration in mules was 123.48±0.61g/L at 1st
week, 126.11±0.13g/L at 2nd week, 122.22±0.17 g/L at 3rd week and
125.85±0.17g/L at 4th week post infection. When diseased and healthy mules
compared on weekly basis, the decrease of haemoglobin concentration was observed
as non significant (P>0.05) among all four weeks. When compared the diseased
horses with diseased mules on weekly basis, haemoglobin concentration was
observed non significant (P>0.05) as shown in fig. 4.16.
Post Infection Weeks
N Horses Mules
Horses Mules Healthy
n=20 Diseased
n=20 Healthy
n=20 Diseased
n=20
1st 40 40 **123.47±0.61a **109.45±0.21a **145.76±0.17a **123.48±0.61b
2nd 40 40 **109.89±0.20d **100.67±0.11c **139.68±0.10b **126.11±0.13a
3rd 40 40 **115.73±0.23c **099.12±0.20d **135.95±0.17c **122.22±0.17c
4th 40 40 **117.24±0.17b **104.61±0.17b **133.97±0.14d **125.85±0.17a
92
Fig 4.16: Week wise comparison of Haemoglobin concentration (g/L) in healthy and diseased horses and mules suffering from strangles. (Mean ± SE)
2- Haematological examination of carrier horses and mules:
For haematological examination of carrier animals, 40 horses (n=20 healthy
horses; n= 20 carrier horses) and 40 mules (n=20 healthy mules; n= 20 carrier mules)
recovered from strangles from Lahore and Sargodha districts of Punjab province of
Pakistan were selected and kept under test upto 8th week of recovery. Parameters like
total white blood cell count, mean segmented neutrophils count, total lymphocytic
count, total monocytic count, total eosinophilic count, total basophilic count, total
erythrocytic count, packed cell volume and haemoglobin concentration were studied.
Total white blood cell count:
Total white blood cell count in carrier horses and mules for strangles is shown
in table 4.31. In horses total WBCs were 14.31±0.19x109/L at 2nd week,
14.07±0.19x109/L at 4th week, 11.69±0.23x109/L at 6th week and 10.88±0.17x109/L
at 8th week of recovery. When compared total WBCs of carrier and healthy horses on
93
weekly basis a significant increase was observed up to 4th week after recovery while
on the 6th and 8th week of recovery this increase was non significant (P>0.05).
In carrier mules total white blood cell count was 13.03±0.2x109/L at 2nd
week, 12.67±0.18x109/L at 4th week, 11.18±0.17x109/L at 6th week and
10.69±0.18x109/L at 8th week of recovery. Similar pattern was observed in carrier
mules as in carrier horses while on comparison of carrier mules with healthy mules
on weekly basis a significant increase was observed up to 4th week of recovery while
on the 6th and 8th of recovery this increase was non significant (P>0.05). It was also
noticed that rise in total white blood cell count was rapid and much more in carrier
horses as compared to carrier mules. When carrier horses compared with carrier
mules, a little difference was observed while total white blood cell count was
significantly less (P<0.05) in normal horses and mules than carrier animals upto 4th
week after recovery as shown in fig. 4.17.
Table 4.31: Total white blood cell count (x109/L) in healthy and carrier horses and mules from strangles. (Mean ± SE)
Mean in a column followed by the same letter were not significantly different at P≤0.05, by Tukey HSD test. * indicates significant difference (p<0.05) among healthy and diseased groups ** indicates non-significant difference (p>0.05) between healthy and diseased group
Weeks After
Recovery
N Horses Mules
Horses Mules Healthy
n=20 Carrier
n=20 Healthy
n=20 Carrier
n=20
2nd 40 40 *11.08±0.13a *14.31±0.19a *10.64±0.12ab *13.03±0.21a
4th 40 40 *10.98±0.17a *14.07±0.19a *10.84±0.09a *12.67±0.18a
6th 40 40 **11.23±0.16a **11.69±0.23b **10.35±0.16b **11.18±0.17b
8th 40 40 **11.20±0.18a **10.88±0.17c **10.26±0.13b **10.69±0.18b
94
Fig 4.17: Week wise comparison of Total white blood cell count (x109/L) in healthy and carrier horses and mules. (Mean ± SE)
Mean Segmented Neutrophilic Count (MSNC):
Data regarding mean segmented neutrophilic count in carrier horses and
carrier mules from strangles is shown in table 4.32. In carrier horses mean segmented
neutrophilic count was 07.13±0.09x109/L at 2nd week, 06.67±0.09x109/L at 4th
week, 04.97±0.13x109/L at 6th week and 04.49±0.07x109/L at 8th week of recovery.
When compared carrier horses with healthy horses on weekly basis significant
increase was observed up to 4th week after recovery while on the 6th and 8th after
recovery this increase was non significant (P>0.05).
In carrier mules mean segmented neutrophilic count was 06.93±0.07x109/L at
2nd week, 05.79±0.11x109/L at 4th week, 05.32±0.07x109/L at 6th week and
04.59±0.08x109/L at 8th week of recovery. When the result of carrier mules
compared with healthy mules on weekly basis significant increase was observed up to
6th week of recovery while on the 8th week of recovery this increase was non
95
significant (P>0.05). The result of carrier mules was different from the result of
carrier horses. When carrier horses compared with carrier mules, no significant
difference was observed (P>0.05) while mean segmented neutrophilic count was
significantly less (P<0.05) in healthy horses and mules than carrier animals upto 4th
week after recovery is shown in fig. 4.18.
Table 4.32: Mean segmented Neutrophils count (x109/L) in healthy and carrier horses and mules from strangles. (Mean ± SE)
Mean in a column followed by the same letter were not significantly different at P≤0.05, by Tukey HSD test. * indicates significant difference (p<0.05) among healthy and diseased groups ** indicates non-significant difference (p>0.05) between healthy and diseased group
Fig 4.18: Week wise comparisons of Neutrophils count (x109/L) in healthy and carrier horses and mules. (Mean ± SE)
Weeks After
Recovery
N Horses Mules
Horses Mules Healthy
n=20 Carrier
n=20 Healthy
n=20 Carrier
n=20
2nd 40 40 *04.37±0.33a *07.13±0.09a *04.71±0.10a *06.93±0.07a
4th 40 40 *04.66±0.40ab *06.67±0.09b *04.62±0.11a *05.79±0.11b
6th 40 40 **04.81±0.52a **04.97±0.13c *04.42±0.10a *05.32±0.07c
8th 40 40 **04.92±0.67a **04.49±0.07d **04.60±0.08a **04.59±0.08d
96
Total Lymphocytic count:
The result of total lymphocytic count in carrier horses and carrier mules for
strangles is shown in table 4.33. Total lymphocytic count in carrier horses was
02.62±0.05x109/L at the end of 2nd week, 03.68±0.09x109/L at the end of 4th week,
03.49±0.13x109/L at the end of 6th week and 02.61±0.04x109/L at the end of 8th
week of recovery. When carrier horses compared with healthy horses there were no
significant increase or decrease (P>0.05) observed among all eight weeks after
recovery.
Table 4.33: Total Lymphocytic count (x109/L) in healthy and carrier horses and mules from strangles. (Mean ± SE)
Mean in a column followed by the same letter were not significantly different at P≤0.05, by Tukey HSD test. * indicates significant difference (p<0.05) among healthy and diseased groups ** indicates non-significant difference (p>0.05) between healthy and diseased group
In mules the values of total lymphocytic count was 03.73±0.06x109/L at 2nd
week, 02.81±0.05x109/L at 4th week, 03.14±0.06x109/L at 6th week and
03.65±0.06x109/L at 8th week of recovery. When compared carrier mules with
healthy mules on weekly basis a significant decrease (P<0.05) was observed in all 8
weeks after recovery and this was opposite to carrier horses. It was also observed that
Weeks After
Recovery
N Horses Mules
Horses Mules Healthy
n=20 Carrier
n=20 Healthy
n=20 Carrier
n=20
2nd 40 40 **02.69±0.07c **02.62±0.05b *05.59±0.04a *03.73±0.06a
4th 40 40 **03.27±0.06b **03.68±0.09a *05.43±0.03a *02.81±0.05c
6th 40 40 **03.65±0.08a **03.49±0.13a *04.77±0.07b *03.14±0.06b
8th 40 40 **03.07±0.05b **02.61±0.04b *04.91±0.07b *03.65±0.06a
97
at the end of 4th week post infection total lymphocytic count was lowest which
significantly increased to the normal values during the subsequent weeks (P<0.05).
When diseased horses compared with diseased mules on weekly basis there was no
significant increase or decrease (P>0.05) observed among all eight weeks after
recovery as shown in fig. 4.19.
Fig 4.19: Week wise comparisons of total lymphocytic count (x109/L) in healthy and carrier horses and mules. (Mean ± SE)
Total Monocytic count (TMC):
Data on total monocytic count in carrier horses and carrier mules from
strangles is shown in table 4.34. Total monocytic count in carrier horses was
00.51±0.019x109/L at the end of 2nd week, 00.44±0.014x109/L at the end of 4th
week, 00.41±0.012x109/L at the end of 6th week and 00.39±0.013x109/L at the end of
8th week of recovery. When compared carrier horses with healthy horses the
observed increase was non significant (P>0.05) among all eight weeks after recovery.
98
Table 4.34: Total Monocyte count (x109/L) in healthy and carrier horses and mules from strangles. (Mean ± SE)
Mean in a column followed by the same letter were not significantly different at P≤0.05, by Tukey HSD test. * indicates significant difference (p<0.05) among healthy and diseased groups ** indicates non-significant difference (p>0.05) between healthy and diseased group
In mules the result of total monocytic count was 00.78±0.02x109/L at 2nd
week, 00.83±0.01x109/L at 4th week, 00.49±0.01x109/L at 6th week and
00.38±0.012x109/L at 8th week of recovery. When compared carrier mules with
healthy mules on weekly basis significant increase was observed up to 4th week of
recovery while on the 6th and 8th of recovery this increase was non significant
(P>0.05). Weekly comparison of carrier horses with carrier mules revealed that
difference of total monocytic count was non significant (P>0.05) among all eight
weeks after recovery as shown in fig. 4.20.
Weeks After
Recovery
N Horses Mules
Horses Mules Healthy
n=20 Carrier
n=20 Healthy
n=20 Carrier
n=20
2nd 40 40 **00.38±0.017a **00.51±0.019a *00.21±0.007ab *00.78±0.024a
4th 40 40 **00.41±0.015a **00.44±0.014b *00.23±0.005a *00.83±0.014a
6th 40 40 **00.37±0.014a **00.41±0.012b **00.19±0.008bc **00.49±0.014b
8th 40 40 **00.38±0.013a **00.39±0.013b **00.17±0.006c **00.38±0.012c
99
Fig 4.20: Week wise comparisons of total Monocytic count (x109/L) in healthy and carrier horses and mules. (Mean ± SE)
Total Eosinophilic count:
The result of total eosinophilic count in carrier horses and carrier mules for
strangles is shown in table 4.35. Total eosinophilic count in carrier horses was
00.47±0.015x109/L at 2nd week, 00.30±0.017x109/L at 4th week, 00.41±0.010x109/L
at 6th week and 00.45±0.017x109/L at 8th week of recovery. When compared carrier
horses with healthy horses the difference of total eosinophilic count was observed non
significant (P>0.05).
100
Table 4.35: Total Eosinophil count (x109/L) in healthy and carrier horses and mules from strangles. (Mean ± SE)
Mean in a column followed by the same letter were not significantly different at P≤0.05, by Tukey HSD test. * indicates significant difference (p<0.05) among healthy and diseased groups ** indicates non-significant difference (p>0.05) between healthy and diseased group
Total eosinophilic count in carrier mules was 00.49±0.014x109/L at 2nd week,
00.51±0.013x109/L at 4th week, 00.47±0.010x109/L at 6th week and
00.40±0.011x109/L at 8th week of recovery. Similarly, the difference of total
eosinophilic count of carrier horses and carrier mules was observed non significant
(P>0.05) among all eight weeks after recovery. Difference between the values of
carrier and healthy mules was also non significant. When compared carrier horses
with carrier mules the difference of total eosinophilic count was observed non
significant (P>0.05) as shown in fig. 4.21.
Weeks After
Recovery
N Horses Mules
Horses Mules Healthy
n=20 Carrier
n=20 Healthy
n=20 Carrier
n=20
2nd 40 40 **00.49±0.017a **00.47±0.015a **00.29±0.011a **00.49±0.014a
4th 40 40 **00.44±0.010b **00.30±0.017c **00.31±0.012a **00.51±0.013a
6th 40 40 **00.48±0.015ab **00.41±0.010b **00.31±0.008a **00.47±0.010a
8th 40 40 **00.47±0.007ab **00.45±0.017ab **00.30±0.007a **00.40±0.011b
101
Fig 4.21: Week wise comparisons of total eosinophilic count (x109/L) in healthy and carrier horses and mules. (Mean ± SE)
Total Basophil count:
Data regarding total basophilic count in carrier horses and carrier mules from
strangles is shown in table 4.36. The result of total basophilic count in carrier horses
was 00.39±0.011x109/L at 2nd week, 00.46±0.016x109/L at 4th week,
00.09±0.015x109/L at 6th week and 00.04±0.005x109/L at 8th week of recovery.
When compared carrier and healthy horses, significant increase was observed up to
4th week of recovery while on the 6th and 8th week of recovery this increase was non
significant (P>0.05).
102
Table 4.36: Total Basophil count (x109/L) in healthy and carrier horses and mules from strangles. (Mean ± SE)
Mean in a column followed by the same letter were not significantly different at P≤0.05, by Tukey HSD test. * indicates significant difference (p<0.05) among healthy and diseased groups ** indicates non-significant difference (p>0.05) between healthy and diseased group
Total basophilic count in carrier mules was 00.19±0.010x109/L at the end of
2nd week, 00.14±0.011x109/L at the end of 4th week, 00.21±0.009x109/L at the end
of 6th week and 00.07±0.012x109/L at the end of 8th week of recovery. When carrier
mules compared with healthy mules, the difference of total basophilic count was
observed non significant (P>0.05). When the result of carrier horses compared with
carrier mules no significant increase (P>0.05) was observed as shown in fig. 4.22.
Weeks After
Recovery
N Horses Mules
Horses Mules Healthy
n=20 Carrier
n=20 Healthy
n=20 Carrier
n=20
2nd 40 40 *00.06±0.005a *00.39±0.011b **00.04±0.004b **00.19±0.010a
4th 40 40 *00.04±0.005b *00.46±0.016a **00.05±0.005ab **00.14±0.011b
6th 40 40 **00.03±0.004bc **00.09±0.015c **00.06±0.005a **00.21±0.009a
8th 40 40 **00.02±0.006c **00.04±0.005d **00.01±0.003c **00.07±0.012c
103
Fig 4.22: Week wise comparisons of total Basophilic count (x109/L) in healthy and carrier horses and mules. (Mean ± SE)
Table 4.30: Total Erythrocytic count:
Data on total erythrocytes count in carrier horses and carrier mules for
strangles is shown in table 4.37. Total erythrocyte count in carrier horses was
5.88±0.15x 1012/L at 2nd week, 5.84±0.09x1012/L at 4th week, 5.76±0.11x 1012/L at
6th week and 5.70±0.06x 1012/L at 8th weeks of recovery. When compared carrier
horses with healthy horses, the decrease of total erythrocytes count was observed non
significant (P>0.05) among all eight weeks of recovery.
Total erythrocytic count in carrier mules was 6.17±0.08x 1012/L at the end of
2nd week, 6.11±0.07x 1012/L at the end of 4th week, 6.50±0.05x 1012/L at the end of
6th week and 6.59±0.08x 1012/L at the end of 8th week of recovery. When carrier
mules compared with healthy mules, the decreases of total erythrocytes count were
observed non significant (P>0.05) as in carrier horses. Similarly when carrier horses
104
compared with carrier mules, the difference of total erythrocytes count was observed
non significant (P>0.05) upto 8th week of recovery as shown in fig. 4.23.
Table 4.37: Erythrocytes count (X 1012/L) in healthy and carrier horses and mules from strangles. (Mean ± SE)
Mean in a column followed by the same letter were not significantly different at P≤0.05, by Tukey HSD test. * indicates significant difference (p<0.05) among healthy and diseased groups ** indicates non-significant difference (p>0.05) between healthy and diseased group
Fig 4.23: Week wise comparisons of Erythrocytes count (X 1012/L) in healthy and carrier horses and mules. (Mean ± SE)
Weeks After
Recovery
N Horses Mules
Horses Mules Healthy
n=20 Carrier
n=20 Healthy
n=20 Carrier
n=20
2nd 40 40 **6.14±0.05a **5.88±0.15a **6.67±0.08a **6.17±0.08b
4th 40 40 **6.08±0.06a **5.84±0.09a **6.59±0.10a **6.11±0.07b
6th 40 40 **6.01±0.05a **5.76±0.11a **6.61±0.09a **6.50±0.05a
8th 40 40 **5.98±0.06a **5.70±0.06a **6.57±0.09a **6.59±0.08a
105
Packed cell volume:
Data regarding packed cell volume in carrier horses and carrier mules from
strangles is shown in table 4.38. In carrier horses packed cell volume was
28.87±0.23% at the 2nd week, 33.08±0.09% at the 4th week, 31.54±0.25% at the 6th
week and 29.22±0.31% at the end of 8th week of recovery. When carrier horses
compared with healthy horses, the decrease of packed cell volume was observed non
significant (P>0.05).
In carrier mules packed cell volume was 29.63±0.23% at the 2nd week,
34.15±0.26% at the 4th week, 32.11±0.51% at the 6th week and 33.57±0.39% at the
end of 8th week of recovery. When compared carrier and healthy mules the difference
of packed cell volume was observed as non significant (P>0.05) in all weeks after
recovery. Similarly the difference of carrier horses with carrier mules packed cell
volume was observed alsonon significant (P>0.05) as shown in fig. 4.24.
Table 4.38: Packed cell volume (%) in healthy and carrier horses and mules from strangles. (Mean ± SE)
Mean in a column followed by the same letter were not significantly different at P≤0.05, by Tukey HSD test. * indicates significant difference (p<0.05) among healthy and diseased groups ** indicates non-significant difference (p>0.05) between healthy and diseased group
Weeks After
Recovery
N Horses Mules
Horses Mules Healthy
n=20 Carrier
n=20 Healthy
n=20 Carrier
n=20
2nd 40 40 **34.98±0.37a **28.87±0.23c **36.82±0.21ab **29.63±0.23c
4th 40 40 **35.75±0.24a **33.08±0.09a **37.10±0.18a **34.15±0.26a
6th 40 40 **33.60±0.33b **31.54±0.25b **36.09±0.26bc **32.11±0.51b
8th 40 40 **35.48±0.29a **29.22±0.31c **35.81±0.26c **33.57±0.39a
106
Fig 4.24: Week wise comparisons of packed cell volume (%) in healthy and carrier horses and mules. (Mean ± SE)
Haemoglobin concentration:
The result of haemoglobin concentration in carrier horses and carrier mules
for strangles is shown in table 4.39. Haemoglobin concentration in carrier horses was
111.32±0.53g/L at 2nd week, 106.87±0.34g/L at 4th week, 105.34±0.29at 6th week
and 110.91±0.60at 8th week of recovery. When carrier and healthy horses compared,
the decrease of haemoglobin concentration was observed non significant (P>0.05). It
was also noted that this decrease of haemoglobin concentration was more in carrier
mules than in carrier horses.
Similarly haemoglobin concentration in carrier mules was 125.53±0.41g/L at
2nd week, 129.20±0.46g/L at 4th week, 130.61±0.42g/L at 6th week and
131.12±0.55g/L at 8th week of recovery. When carrier and healthy mules compared
the decrease of haemoglobin concentration was observed non significant (P>0.05)
during all weeks after recovery. When carrier horses compared with carrier mules the
107
haemoglobin concentration was observed non significant (P>0.05) as shown in fig.
4.25.
Table 4.39: Haemoglobin concentration (g/L) in healthy and carrier horses and
mules from strangles. (Mean ± SE)
Mean in a column followed by the same letter were not significantly different at P≤0.05, by Tukey HSD test. * indicates significant difference (p<0.05) among healthy and diseased groups ** indicates non-significant difference (p>0.05) between healthy and diseased group
Fig 4.25: Week wise comparisons of Hb concentration (g/L) in healthy and carrier horses and mules. (Mean ± SE)
Weeks After
Recovery
N Horses Mules
Horses Mules Healthy
n=20 Carrier n=20
Healthy n=20
Carrier n=20
2nd 40 40 **124.51±0.37a **111.32±0.53a **141.81±0.79a **125.53±0.41c
4th 40 40 **115.43±0.84c **106.87±0.34b **137.73±0.40bc **129.20±0.46b
6th 40 40 **110.89±0.52d **105.34±0.29b **139.48±0.59b **130.61±0.42ab
8th 40 40 **119.37±0.47b **110.91±0.60a **136.22±0.26c **131.12±0.55a
108
Protein Analysis:
For protein analysis 40 equids (n=20 horses, n= 20 mules) under 5 years of
age suffering from natural outbreak of strangles from Lahore and Sargodha districts
of Punjab province of Pakistan were selected for recording of observations.
Parameters studied included, total serum protein, serum albumin, serum globulin and
fibrinogen.
Total Serum Proteins: Total serum protein values in horses and mules suffering from strangles are
shown in table 4.40. In horses total serum protein values 82.7±3.3g/L under one year
of age, 80.9±5.7g/L between 1-2 year of age, 79.3±6.4g/L between 2-3 year of age,
78.8±5.3g/L between 3-4 year of age and78.5±4.7g/L between 4-5 year of age were
observed. When compared total serum protein values of diseased and healthy horses
on age basis, a significant increase (P<0.05) was observed in all age groups. Among
all age groups total serum protein values were according to the following order 5th <
4th < 3rd< 2nd < 1st week. It was also observed that total serum protein value was
highest in group which was under one year of age, it significantly decreased to the
normal during the subsequent weeks (P<0.05).
In mules total serum protein values 81.5±5.3g/L under one year of age,
83.1±6.3g/L between 1-2 years of age, 80.1±6.4g/L between 2-3 years of age,
79.3±5.5g/L between 3-4 years of age and78.4±4.8g/L between 4-5 years of age were
observed. When compared total serum protein values of diseased and healthy mules
of different age groups, a significant increase (P<0.05) was observed in all age
groups. Among all age groups total serum protein values were according to the
109
following order 5th < 4th < 3rd< 2nd < 1st week. It was also observed that total serum
protein values were highest in group having age under one year which significantly
decreased to the normal in the subsequent age groups (P<0.05).When diseased horses
compared with diseased mules, no significant difference was observed while total
serum protein values were significantly less (P<0.05) in healthy horses and mules
than diseased as shown in fig. 4.26.
Table 4.40: Total serum protein values (g/L) of healthy and diseased horses and mules suffered from strangles. (Mean ± SD)
Mean in a row followed by the same letter were not significantly different at P≤0.05, by paired T test.
Fig 4.26: Age wise comparison of total serum protein values (g/L) of healthy and diseased horses and mules suffered from strangles. (Mean ± SD)
Groups N
Horses Mules Horses Mules
Healthy Diseased Healthy Diseased Healthy Diseased Healthy Diseased
< 1year 10 10 10 10 64.9a ±3.4
82.7b ±3.3
65.8a ± 3.5
81.5b ± 5.3
1-2 year
10 10 10 10 66.5a ±3.1
80.9b ±5.7
68.0a ± 3.8
83.1b ± 6.3
2-3 year
10 10 10 10 62.7a ±1.2
79.3b ±6.4
67.3a ± 4.4
80.1b ±6.4
3-4 year
10 10 10 06 63.6a ±5.8
78.8b ±5.3
64.7a ± 4.8
79.3b ±5.5
4-5 year
10 10 10 03 64.1a ±4.1
78.5b ±4.7
65.0a ± 4.4
78.4b ±4.8
110
Serum Albumin:
Data on serum albumin in horses and mules suffering from strangles are
shown in table 4.41. In horses serum albumin values 25.4±2.9g/L under one year of
age, 27.1±3.9g/L between 1-2 years of age, 24.8±4.4g/L between 2-3 years of age,
26.2±4.5g/L between 3-4 years of age and28.4±5.4g/L between 4-5 years of age were
observed. When compared serum albumin values of diseased and healthy horses on
age basis significant decrease (P<0.05) was observed in all age groups.
In mules serum albumin values 26.3±4.2g/L under one year of age,
26.1±4.2g/L between 1-2 years of age, 27.5±4.0g/L between 2-3 years of age,
28.3±5.1g/L between 3-4 years of age and28.7±5.4g/L between 4-5 years of age were
observed. Age based comparison of serum albumin values of diseased and healthy
horses revealed a significant decrease (P<0.05).
Table 4.41: Serum albumin values (g/L) of healthy and diseased horses and mules suffered from strangles. (Mean ± SD)
Mean in a row followed by the same letter were not significantly different at P≤0.05, by paired T test.
Groups N
Horses Mules Horses Mules
Healthy Diseased Healthy Diseased Healthy Diseased Healthy Diseased
< 1year 10 10 10 10 33.8a
±2.2
25.4b
±2.9
34.4a
± 2.6
26.3b
±4.2
1-2 year 10 10 10 10 36.3a
±2.4
27.1b
±3.9
34.3a
± 3.0
26.1b
±4.2
2-3 year 10 10 10 10 34.7a
±4.4
24.8b
±4.4
36.1a
± 5.2
27.5b
±4.0
3-4 year 10 10 10 06 35.3a
±5.3
26.2b
±4.5
35.2a
± 5.1
28.3b
±5.1
4-5 year 10 10 10 03 34.5a
±4.1
28.4b
±5.4
36.5a
± 4.9
28.7b
±5.4
111
Fig 4.27: Age wise comparison of serum albumin values (g/L) of healthy and diseased horses and mules suffered from strangles. (Mean ± SD)
When diseased horses compared with diseased mules, no significant
difference was observed while serum albumin values were significantly less (P<0.05)
in diseased horses and mules than healthy as shown in fig. 4.27.
Serum Globulin:
Serum globulin in horses and mules suffering from strangles are shown in
table 4.42. In horses serum globulin values 37.1±3.4g/L under one year of age,
36.7±4.5g/L between 1-2 years of age, 35.2±4.0g/L between 2-3 years of age,
36.7±2.7g/L between 3-4 years of age and 32.9±1.9g/L between 4-5 years of age
were observed. When compared serum albumin values of diseased and healthy horses
on age basis, a significant increase (P<0.05) was observed in group having age under
one year, while rest of the age groups showed a non significant increase.
112
Table 4.42: Serum globulin values (g/L) of healthy and diseased horses and mules suffered from strangles. (Mean ± SD)
Mean in a row followed by the same letter were not significantly different at P≤0.05, by paired T test.
Fig 4.28: Age wise comparison of serum globulin values (g/L) of healthy and diseased horses and mules suffered from strangles. (Mean ± SD)
Groups N
Horses Mules Horses Mules
Healthy Diseased Healthy Diseased Healthy Diseased Healthy Diseased
< 1year 10 10 10 10 32.9a
±1.8
37.1b
±3.4
33.7a
± 1.2
36.2b
±3.8
1-2 year 10 10 10 10 34.5a
±3.0
36.7a
±4.5
33.6a
± 2.9
37.3b
±4.3
2-3 year 10 10 10 10 33.2a
±6.5
35.2a
±4.0
33.9a
± 5.6
35.3a
±4.0
3-4 year 10 10 10 06 34.4a
±3.2
36.7a
±2.7
33.5a
± 2.7
34.9a
±3.5
4-5 year 10 10 10 03 33.7a
±4.4
32.9a
±1.9
33.8a
± 4.5
33.1a
±1.9
113
In mules serum albumin values 36.2±3.8g/L under one year of age,
37.3.1±4.3g/L between 1-2 years of age, 35.3±4.0g/L between 2-3 years of age,
34.9±3.5g/L between 3-4 years of age and33.1±1.9g/L between 4-5 years of age were
observed. When compared serum albumin values of diseased and healthy horses on
age basis, a significant increase (P<0.05) was observed in group having age from one
to two years. It was different from the horses where significant increase was observed
only in the group which was under one year of age. When diseased horses compared
with diseased mules, no significant difference was observed while serum globulin
values were increased but not significantly in diseased horses and mules than healthy
as shown in fig. 4.28.
Fibrinogen values:
Data on fibrinogen values in horses and mules suffering from strangles are
shown in table 4.43. In horses fibrinogen values 7.2±2.3g/L under one year of age,
6.7±2.4g/L between 1-2 years of age, 6.3±2.2g/L between 2-3 years of age,
5.7±1.0g/L between 3-4 years of age and 5.1±0.7g/L between 4-5 years of age were
observed. Among all age groups fibrinogen values were according to the following
order 5th < 4th < 3rd< 2nd < 1st week. It was also observed that fibrinogen values
were highest in under one year of age group which significantly decreased to the
normal values in the subsequent age groups (P<0.05). When compared fibrinogen
values of diseased and healthy horses on age basis significant increase (P<0.05) was
observed in all age groups that indicate significance of hyperfibrinegemia for
diagnosis of diseased.
114
In mules fibrinogen values 6.5±3.1g/L under one year of age, 7.0±2.6g/L
between 1-2 years of age, 6.0±2.3g/L between 2-3 years of age, 5.0±1.4g/L between
3-4 years of age and 4.5±1.2g/L between 4-5 years of age were observed. Similarly in
mules fibrinogen values were according to the following order 5th < 4th < 3rd< 2nd <
1st week in all age groups. It was also observed that fibrinogen values were highest
under one year of age group which significantly decreased to the normal in the
subsequent age groups (P<0.05).When compared fibrinogen values of diseased and
healthy horses on age basis significant increase (P<0.05) was observed in all age
groups that was similar to horses. When diseased horses compared with diseased
mules, no significant difference was observed while fibrinogen values were
significantly increased (P<0.05) in diseased horses and mules than healthy as shown
in fig. 4.29.
Table 4.43: Fibrinogen values (g/L) of healthy and diseased horses and mules suffered from strangles. (Mean ± SD)
Mean in a row followed by the same letter were not significantly different at P≤0.05, by paired T test.
Groups N
Horses Mules Horses Mules
Healthy Diseased Healthy Diseased Healthy Diseased Healthy Diseased
< 1year 10 10 10 10 03.4a
±1.0
07.2b
±2.3
03.3a
±1.0
06.5b
±3.1
1-2 year 10 10 10 10 03.8a
±1.0
06.7b
±2.4
03.6a
±1.2
07.0b
±2.6
2-3 year 10 10 10 10 03.5a
±1.1
06.3b
±2.2
03.9a
±1.5
06.0b
±2.3
3-4 year 10 10 10 06 03.7a
±0.7
05.7b
±1.0
03.6a
±0.9
05.0b
±1.4
4-5 year 10 10 10 03 03.6a
±1.4
05.1b
±0.7
03.8a
±1.4
04.5b
±1.2
115
Fig 4.29: Age wise comparison of fibrinogen values (g/L) of healthy and diseased horses and mules suffered from strangles. (Mean ± SD)
THERAPEUTIC TRIALS:
In-vitro Antibiotic sensitivity test of horses:
Data on twenty randomly selected β hemolytic colonies of horses
subjected to sensitivity of ten antibiotics is shown in table 4.44. It was observed that
higher number of field samples of S. equi were sensitive to Procaine penicillin
followed by ceftiofur Na, cephradine, erythromycin, ampicillin, tetracycline,
chloramphenicol, sulfamethoxazole, trimethoprim +sulphadiazine and gentamycin,
respectively. It is concluded from the result of present study that field isolate of S.
equi is still more sensitive to Procaine penicillin, ceftiofur Na, cephradine and
erythromycin as compared to other antibiotics.
116
Table 4.44: In-vitro Antibiotic sensitivity against S. equi in horses
ZI= zone of inhibition
In-vitro Antibiotic sensitivity test of mules:
Data regarding, twenty randomly selected β hemolytic colonies of S. Equi of
mules subjected to sensitivity of ten antibiotics is shown in table 4.45. In the present
study it was also observed that higher number of field isolates of S. equi were
sensitive to Procaine penicillin followed by ceftiofur Na, erythromycin, cephradine,
ampicillin, tetracycline, chloramphenicol, sulfamethoxazole, trimethoprim +
sulphadiazine and gentamycin that was similar to horses.
Antibiotics Sensitive Discs No. of Horses
Remarks Sensitive Intermediate Resistant
No. of isolates
ZI (mm)
No. of isolates
ZI (mm)
No. of isolates
ZI (mm)
Procaine Penicillin 20 19 >30 01 28-29 - <28
Ceftiofur Na 20 18 >27 01 25-26 01 <25
Cephradine 20 16 >25 02 23-24 02 <23
Erythromycin 20 17 >22 02 20-21 01 <20
Tetracycline 20 12 >16 06 14-15 02 <14
Sulfamethoxazole 20 13 >14 04 12-13 03 <12
Trimethoprim+sulphadiazine 20 12 >12 03 10-11 05 <10
Chloramphenicol 20 13 >15 05 12-14 02 <12
Ampicillin 20 14 >19 04 15-18 02 <15
Gentamycin 20 12 >10 06 7-9 02 <7
117
Table 4.45: In-vitro Antibiotic sensitivity against S. equi in mules
Antibiotics Sensitive Discs No. of Mules
Remarks Sensitive Intermediate Resistant
No. of isolates
ZI (mm)
No. of isolates
ZI (mm)
No. of isolates
ZI (mm)
Procaine Penicillin 20 19 >30 01 28-29 - <28
Ceftiofur Na 20 18 >27 02 25-26 - <25
Cephradine 20 15 >25 04 23-24 01 <23
Erythromycin 20 16 >22 02 20-21 02 <20
Tetracycline 20 13 >16 04 14-15 03 <14
Sulfamethoxazole 20 15 >14 03 12-13 02 <12
Trimethoprim+sulphadiazine 20 14 >12 04 10-11 02 <10
Chloramphenicol 20 12 >15 04 12-14 04 <12
Ampicillin 20 15 >19 02 15-18 03 <15
Gentamycin 20 14 >10 05 7-9 01 <7
ZI= zone of inhibition
In-vivo Antibiotic sensitivity test of horses:
Out of ten antibiotics used in-vitro antibiotic sensitivity test against S. equi,
four top ranking antibiotics were selected for in-vivo trials in horses.
Group A:
The ten horses (n=5 without abscess, n=5 with abscess) of group A were
treated with rank 1 antibiotic i.e. Procaine Penicillin is shown in table 4.46. The
efficacy of Procaine Penicillin was measured on the basis of disappearance of clinical
118
signs. Response to the antibiotic in the horses without abscess was excellent while the
horses with abscess exhibited a poor response.
Group B:
The ten horses (n=5 without abscess, n=5 with abscess) of group B were
treated with rank 2 antibiotic i.e. ceftiofur Na is shown in table 4.46. The criteria for
measuring the efficacy of ceftiofur Na was same as in Group A. The results in horses
without abscess were very good while the abscessed horses showed poor results.
Group C:
The ten horses (n=5 without abscess, n=5 with abscess) of group C were
treated with rank 3 antibiotic i.e. cephradine is shown in table 4.46. The efficacy of
cephradine was measured on similar basis as in above mentioned groups. The results
showed good response in horses without abscess whereas the response was poor in
horses with abscess.
Group D:
The ten horses (n=5 without abscess, n=5 with abscess) of group D were
treated with rank 4 antibiotic i.e. erythromycin is shown in table 4.46. The efficacy of
erythromycin was measured on the basis of disappearance of clinical signs in horses.
Like the other groups, the results in horses without abscess were good while the
horses with abscess showed poor results.
119
Table 4.46: In-vivo Antibiotic sensitivity against S. equi in horses
Groups Antibiotics used No. of horses (n=10)
Without abscess(n=5)
With abscess(n=5)
A Procaine Penicillin Excellent poor
B Ceftiofur Na Very good poor
C Cephradine Good poor
D Erythromycin Good poor
In-vivo Antibiotic sensitivity test of mules:
Similarly in mules out of ten antibiotics, four top ranked antibiotics were
selected on the basis of in-vitro antibiotic sensitivity test and used as in-vivo trials in
mules.
Group A:
A group of ten mules comprised of n=5 without abscess, n=5 with abscess of
group A were treated with rank 1 antibiotic i.e. Procaine Penicillin is shown in table
4.47. The efficacy of procaine penicillin was measured on the basis of disappearance
of clinical signs in the mules. The results of present study showed excellent response
in mules without abscess while the response was poor in mules with abscess.
Group B:
A group of ten mules comprised of n=5 without abscess, n=5 with abscess of
group B were treated with rank 2 antibiotic i.e. Ceftiofur Na, results are shown in
table 4.47. The efficacy of Ceftiofur Na was measured on similar basis as in group A.
120
The results in mules without abscess were very good while in mules with abscess
were poor.
Group C:
A group of ten mules (n=5 without abscess, n=5 with abscess) of group C
were treated with rank 3 antibiotic i.e. Cephradine, results are shown in table 4.47.
The criteria for measuring the efficacy of antibiotics was the disappearance of clinical
signs and results obtained in mules without abscess were good while poor in
abscessed mules.
Group D:
A group of ten mules (n=5 without abscess, n=5 with abscess) of group D
were treated with rank 4 antibiotic i.e. erythromycin, results are shown in table 4.47.
The efficacy of erythromycin was measured on the basis of disappearance of clinical
signs in the mules and similar results were obtained as in Group C.
Table 4.47: In-vivo Antibiotic sensitivity against S. equi mules
Groups Antibiotics used No. of mules (n=10)
Without abscess(n=5)
With abscess(n=5)
A Procaine Penicillin Very good poor
B Ceftiofur Na Excellent poor
C Cephradine Good poor
D Erythromycin Good poor
121
It is concluded from the result of present study that treatment should be started
as early as possible before the appearance of abscess in both horses and mules.
In-vitro Disinfectant Trial:
In-vitro testing of S. equi with various disinfectants was performed by using
the Phenol Co-efficient Test to determine susceptibility of bacteria. For that purpose,
four disinfectants Dettol, Povidone iodine, 0.6% H2SO4 and Bleach were used in the
study.
In-vitro Disinfectant trial of Phenol:
Data regarding the growth of S. equi after using phenol as disinfectant in
different interval of time is shown in table 4.48. In-vitro disinfectant trial of Phenol
was made to compare with other disinfectants used in the study. For that purpose six
serial dilutions of phenol were made in separate tubes as follows 1:50, 1:100, 1:150,
1:200, 1:250 and 1:300. Samples from these tubes were then plated on blood agar
plates after 5, 10 and 15 minutes. The highest dilution of phenol was recorded as
1:200.
Table 4.48: In-Vitro efficacy of Phenol as disinfectant against S. equi
Phenol
Dilution 5 minutes 10 minutes 15minutes
1 to 50 No growth No growth No growth
1 to 100 Growth No growth No growth
1 to 150 Growth No growth No growth
1 to 200 Growth No growth No growth
1 to 250 Growth Growth No growth
1 to 300 Growth Growth No growth
122
In-vitro Disinfectant trial of Dettol:
The result on the growth of S. equi after using dettol as disinfectant is given in
table 4.49. For in-vitro disinfectant trial of dettol, six serial dilutions were made as
follows 1:50, 1:100, 1:150, 1:200, 1:250 and 1:300 in separate tubes. The time used to
take the samples from the tubes and to check the growth on the blood agar was same
as in phenol. After that highest dilution of dettol was recorded as 1:100 while phenol
was 1:200. The phenol coefficient was calculated by dividing the highest dilution of
dettol over highest dilution of phenol.
Table 4.49: In-Vitro efficacy of Dettol as disinfectant against S. equi
Dettol
Dilution 5 minutes 10 minutes 15minutes
1 to 50 No growth No growth No growth
1 to 100 Growth No growth No growth
1 to 150 Growth Growth No growth
1 to 200 Growth Growth Growth
1 to 250 Growth Growth Growth
1 to 300 Growth Growth Growth
Phenol Coefficient=100/200=0.5
In-vitro Disinfectant trial of Povidone Iodine:
Data on the growth of S. equi after using povidone iodine as disinfectant is
given in table 4.50. For povidone iodine in-vitro disinfectant trial six serial dilutions
123
were made as were in phenol and dettol. The time used to take samples was 5, 10 and
15 minutes. The highest dilution of povidone iodine was recorded as 1:250 and the
phenol coefficient was calculated as 1.25.
Table 4.50: In-Vitro efficacy of Povidone Iodine as disinfectant against S. equi
Povidone
Iodine
Dilution 5 minutes 10 minutes 15minutes
1 to 50 No growth No growth No growth
1 to 100 Growth No growth No growth
1 to 150 Growth No growth No growth
1 to 200 Growth No growth No growth
1 to 250 Growth No growth No growth
1 to 300 Growth Growth No growth
Phenol Coefficient=250/200=1.25
In-vitro Disinfectant trial of 0.6%H2SO4:
The result on the growth of S. equi after using 0.6% H2SO4 as disinfectant is
given in table 4.51. For in-vitro disinfectant trial of 0.6% H2SO4 six serial dilutions
were made as in above disinfectants. The time used to take samples from the dilutions
was same as in above disinfectants. The highest dilution was recorded as 1:200 and
phenol coefficient was 1.00.
124
Table 4.51: In-Vitro efficacy of 0.6% Sulfuric acid as disinfectant against S. equi
Phenol Coefficient=200/200=1.00
In-vitro Disinfectant trial of Bleach:
The results on the growth of S. equi after using bleach as disinfectant are
given in table 4.52. For in-vitro disinfectant trial of bleach serial dilutions were made
as in above disinfectants. Samples from different dilutions were taken after the same
time as in other disinfectants used in the study. Highest dilution was 1:150. The
phenol coefficient was calculated 0.75.
Table 4.52: In-Vitro efficacy of Bleach as disinfectant against S. equi
Bleach
Dilution 5 minutes 10 minutes 15minutes
1 to 50 No growth No growth No growth
1 to 100 Growth No growth No growth
1 to 150 Growth No growth No growth
1 to 200 Growth Growth No growth
1 to 250 Growth Growth No growth
1 to 300 Growth Growth No growth
Phenol Coefficient=150/200=0.75
0.6%H2SO4
Dilution 5 minutes 10 minutes 15minutes
1 to 50 No growth No growth No growth
1 to 100 Growth No growth No growth
1 to 150 Growth No growth No growth
1 to 200 Growth No growth No growth
1 to 250 Growth Growth No growth
1 to 300 Growth Growth No growth
125
Comparison of Disinfectants:
When compared all four disinfectants used against S. equi in the present study
with phenol by using the Phenol Co-efficient Test, povidone iodine was found to be
the best one because its phenol coefficient was 1.25 that was greater than phenol 1.00
while 0.6% H2SO4 showed similar phenol coefficient as that of phenol. The dettol and
bleach showed phenol coefficient lesser than phenol as shown in table 4.53. Therefore
use of povidone iodine and 0.6% H2SO4 on priority basis against S. equi is
recommended as compared to other disinfectants.
Table 4.53: Overall comparison of Different Disinfectants used against S. equi
Disinfectants Highest Dilution of disinfectant
Highest dilution of
Phenol
Phenol Coefficient
Remarks
Dettol 1:100 1:200 100/200=0.5 Fair
Povidone Iodine 1:250 1:200 250/200=1.25 Very good
Bleach 1:150 1:200 150/200=0.75 Satisfactory
0.6%H2SO4 1:200 1:200 200/200=1.00 Good
126
Chapter-05
DISCUSSION
EPIDEMIOLOGICAL STUDIES:
PREVALENCE OF HORSES AND MULES:
Data on the overall prevalence of S. equi on the basis of culture in horses and
mules is shown in table 4.1 and 4.7. Out of 250 horses and 250 mules, 113(45.2%)
horses and 99 (39.6%) mules tested positive for S. equi. Minett, 1944 and Peat, 1945
reported the incidence of strangles in both horses and mules at Remount depot Mona
and Sargodha, Pakistan. The number of S. equi isolates were significantly higher
(P<0.05) in pus samples collected from sub-mandibular lymph nodes as compared to
nasal discharge samples. The difference in number was found significant (P<0.05)
among horses and mules of different age groups. The highest prevalence of strangles
was recorded in horses and mules aging less than 2 years as compared to those having
age more than 2 years. Similar findings were reported by Fallon, 1969. He reported
that the disease is more prevalent in young animals, especially in populations with a
prior history of strangles outbreaks. Results of our study also correlate with the
findings of Timoney, 1993 who also reported that horses of all ages may be affected,
but the disease is most common and most severe in young horses. It is therefore
mostly prevalent on breeding farms. Outbreaks are often initiated by the introduction
of an animal to the farm that is either incubating the disease or is still shedding the
organism during the recovery phase. Sweeny, 1987 and Piche, 1984 observed that
yearlings and young adults are most at risk, followed by weanlings and then adults.
127
Typically, yearlings are most severely affected with a longer duration of clinical signs
(Piche, 1984). During the breeding season, nursing mares brought of suckling foals
may introduce S. equi in this manner. Findings of our study are also in line with the
findings of Sweeny, 1990 who reported that infection occurs primarily in 1 to 5 years
old horses, but is not restricted to age groups. As regards incidence of strangles from
the end of January to the beginning of May was found to be the highest (2.6%) in
foals of 9 months–2 years compared to adult mules 2–5 years old, confirming that
once an animal is infected with strangles it attains lifelong immunity (Walker and
Timoney, 1998). Our results were also broadly consistent with the findings of Ashraf
(2000) who reported 33.2% strangles in Pakistan in mules less than 2 year of age and
35.4% in mules of more than 2 year of age. Similarly, results of the present study are
in agreement with the findings of Manzoor et al. 2008 who reported 54% infection in
foals in Punjab, Pakistan. Our findings are also in agreement with the results of
Sweeney et al., 1989 who found rates of S. equi infections of the upper respiratory
tract and lymph nodes (strangles) in horses to be 47.5% for 1-year-old horses, and
37.5% for foals. S. equi was isolated from nasal, pharyngeal, or lymph node
specimens in 31 (60.8%) of 51 sick horses. Our results also correlate with the study of
Hamlen et al., 1994 who reported that foals were highly susceptible to developing
strangles following S. equi exposure as shown by attack rates of 86% (19/22) and
91% (10/11) respectively.
In the present study the prevalence of strangles in horses and mules were also
calculated round the year, it was found to be the highest during the months of
February, March, April and May, while few cases were seen during the months of
128
January, June and July and no cases were seen during rest of the months. The
significant difference was observed (p<0.05) among different months of year.
Similarly when compared the prevalence of strangles in different seasons of Pakistan,
the highest prevalence rate was recorded during the spring months. Our study
correlates with the findings of Manzoor et al., 2008 who recorded incidence of
strangles in foals of 9 months to 2 years of age and it was found to be the highest
during the spring season (Mid of January to Start of May).
Data regarding the overall prevalence of strangles on the basis of Polymerase
chain reaction of S. equi in equines is also shown in table 4.4 and 4.10. Out of Equine
groups of 250 horses and mules each, 122(48.8%) horses and 113(45.2%) mules
tested positive for S. equi. On comparison of the prevalence rate on the basis of PCR
and culture, (nasal swabs or pus samples from affected submandibular lymph nodes)
the sensitivity of Polymerase chain reaction appeared to be much greater than culture.
Laboratory experimentation has demonstrated a detection sensitivity of 10 or fewer
colony forming units (Timoney and Artiushin, 1997). The culture along with PCR is
best techniques for diagnosing S. equi. Culture is of value because it definitely
establishes infection and can conviently be performed on the same samples used for
PCR. Since PCR can be completed in four to five hours it should be an effective tool
in the management of outbreaks and in screening equines before and after
transportation (Timoney and Artiushin, 1997). Newton et al., 2000 reported that PCR
has been developed to detect the DNA sequence of the S. equi SeM gene, and can be
used to confirm the diagnosis of strangles within hours of sample submission.
Because this test does not differentiate between dead and live bacteria, a positive test
129
may not correlate with active infection; therefore, a positive culture may be necessary
to confirm the diagnosis. Timoney and Artiushin, 1997 also reported that PCR is
approximately three times more sensitive than the culture in detecting S. equi. The
relative insensitivity of the culture may be due to an inadequate number of collected
organisms, overgrowth of contaminants, or slow growth.
MORTALITY RATE:
Data regarding the mortality rate in horses and mules are shown in table 4.13
and 4.14. In horses and mules the mortality was recorded 1.64% and 0.88%,
respectively under 5 years of age. The non significant (P>0.05) difference was
observed in mortality rate of affected equines among different age groups. From the
present study it was concluded that the severity of disease is greater in animals less
than 2 years of age as compared to over two year of age. The results of the present
study were also in line with the findings of Higgins and Snyder (2006) who reported
mortality rates between 1-5%. Mortality is generally low (1-2%) in uncomplicated
cases (McAllister, 1982; Radostits et al., 2000; Bryans and Moore, 1972; McGee,
1969). Radostits et al. (2000) also reported that most deaths are due to the
dissemination of organisms to the other organs. Various research workers reported
wide range of mortality in equines due to this disease which include Minett, 1944
who recorded mortality as 5% at Mona and 6.1% at Sargodha in horses and 1% in
mules at both the depots. Similarly Wilson, 1988 recorded mortality rates as 1-5%. A
2.6% mortality was recorded by Sweeny et al., 1989 whereas 1-2%, 4.4% and zero
percent mortality was recorded by Clabough, 1987, Vukovic, 1961 and Zedeh et al.,
1992 respectively.
130
VARIATIONS IN SeM, SzPSe & Se 18.9 PROTEINS OF S.equi:
SeM:
PCR of the SeM genes of 24 selected isolates of S. equi and the US prototype
strain CF32 generated products of 1812 bp whereas resulting DNA sequences
encoded 19 different SeM alleles including numbers 71-76 not previously included in
the database (www.pubmlst.org/szooepidemicus/) (Table 4.16). The results confirmed
the great variability of the N-terminus of SeM noted in previous studies (Anzai et al.,
2005; Kelly et al., 2006). Residues most frequently subject to substitution were
located at position 58, 63, 108 and 143. Single nucleotide polymorphisms (SNPs) in
SeM were found at 93 loci and totaled 181. Fifty-eight of these were non-
synonymous, that is, were mutations resulting in amino acid replacements in SeM.
Non-synonymous SNPs were 15.7 times more frequent in the N-terminal region
(positions 114 to 629) than in the remainder of the SeM sequence (Table 4.18). These
results are consistent with a previous estimate (3.054) of the ratio of non-synonymous
to synonymous amino acid substitutions (dN/dS) in the SeM N-terminus (Kelly et al.,
2006). Discovery of non-synonymous substitutions distal to the N-terminus indicates
this region is also under diversifying selection pressure albeit of lower magnitude
than in the N-terminus. Substitution of amino acids in this region has been shown to
alter a conformational epitope, and is more likely due to immune selection of mucosal
IgA or local T cell rather than serum IgG responses (Timoney et al., 2009). Since
emergence of variants has been detected in chronically infected guttural pouches and
cranial sinuses, alteration in conformation of the N-terminus may serve as a
mechanism of immune escape in these niches as well as implicating an unknown
131
interaction of the N-terminus of SeM with mucosal IgA or T-cell responses that
reduces survival of the bacterium. Fibrinogen binding or opsonization of S.equi by
SeM specific antibody are not significantly affected by N1 terminal variation
(Timoney et al., 2009). The survival value 1 of allelic variation seems to be restricted
to the mucosal compartment, since horses with chronic guttural pouch infections do
not experience recurrence of clinical strangles caused by the newly emerging SeM
allelic variants. In these cases, the acquired systemic immune response continues to
protect the animal’s regional lymph nodes and tonsils. Phylogenies generated by
neighborhood joining indicate that SeM allele 71 identified in isolates S24 and L32
from Pakistan was the most distantly related allele of the 25 isolates in the study. This
is explained by the remote and isolated Pakistani equid population, which
hypothetically favors preservation and continued divergence of a specific SeM allele.
Other newly identified alleles (72 - 76) in N. America isolates showed 96 - 99%
similarity with alleles 62, 59, 57 and 37 in the SeM database.
SzPSe:
With one exception, sequence analysis of the SzPSe genes provided no
instances of variation. The single exception, Australian isolate 181, had a deletion of
one PEPK repeat remarkably, although 92 SNPs were found at 48 loci in SzPSe of the
25 S. equi isolates including S. equi CF32, no SNPs encoding non-synonymous
substitutions were found (Table 4.19). In the horses, SzPSe elicits strong serum IgG
and mucosal IgA responses during recovery from strangles and so is exposed to
immune selection pressures similar to those exerted on SeM (Timoney et al., 2007)
although the results indicate that selection is purifying and not diversifying, moreover
132
survival of S. equi in the horses requires conservation of SzPSe. This may be related
to a binding function of SzP to receptors in the crypts of the lingual and palatine
tonsils, sites of entry of S. equi (Kumar et al., 2007). The epitope responsible for
binding of SzP requires a specific conformation (Fan et al., 2008). Unlike S. equi, SzP
alleles in S. zooepidemicus are highly variable and are the basis for the Moore and
Bryans typing scheme (Moore and Bryans, 1969). The SzP family of proteins shows 2
forms of N-terminal variation, N1 and N2, at least 5 variants of a central non–alpha-
helical hypervariable region, HV1 to 5 and a variable number of carboxyterminal
PEPK repeats (Walker and Timoney, 1998). Thus, there is evidence of a high rate of
recombination in the SzP gene.
Moreover, the HV region appears to have been horizontally acquired since its
G-C % (38.3) differs significantly (p<0.01) from that of the remaining SzPSe
sequence (47.0). Recombination and the presence of exogenous DNA sequence are
factors that would favor occurrence of SNPs. The biological/immunological
significance of this variation is not understood, but does not appear to involve
opsonogenic epitopes. Future work might logically address the effect of variation on
the conformational adhesion epitope on host cell specificity.
Se18.9:
The action of highly conserved Se18.9 is anti-phagocytic in nature. It is
accomplished by binding complement control factor H and reducing the bactericidal
activity of neutrophils by an unknown mechanism (Tiwari et al., 2007). Uniquely
expressed by S. equi, Se18.9 binds to tonsillar epithelium and stimulates strong
mucosal IgA and serum IgG responses. These antibodies neutralize the bactericidal
133
activity of Se18.9 of neutrophils. Since this protein has a proven virulence function in
common with SeM, it is logically a target of immune selection pressure yet only 2
SNP loci have been found in the DNA sequences of the Se18.9 genes in 25 isolates.
The unexpected absence of variants of Se18.9 in a population of SeM allelic variants
of S. equi argues either for an immutable and essential structure or virulence function
that is minor compared to that of SeM. It might also be argued that as a secreted
protein, Se18.9 might have less survival value for Se than a protein anchored on its
surface. The much lower frequency of SNP loci in Se18.9 compared to SeM and
SzPSe is unexplained.
Mutation plays an important role in a pathogen’s accommodation to adaptive
immune responses, which in turn favors its persistence and transmission to new hosts
(Aguileta et al., 2009). The remarkable genetic and phenotypic conservation of the
almost colonial S. equi compared to the closely related S. zooepidemicus is consistent
with its adaptation to a specific host. Variations in virulence proteins such as SzPSe
and Se18.9 may reduce its fitness to infect and replicate and so colonies of S. equi
with mutations that affect these proteins do not survive. In contrast, changes in the N7
terminal sequence of SeM provide a survival advantage in niches such as the guttural
pouch or cranial sinuses where mucosal responses dominate the host – parasite
interaction.
STUDY OF CARRIER STATUS OF HORSES & MULES:
Data regarding carrier state of naturally infected horses and mules are shown
in table 4.20 and 4.21. Out of 122 positive horses and 113 positive mules for
strangles, 20 horses and 20 mules (10 < 2 year and 10 between 2 and 5 years of age)
134
remaining positive after one week of infection were monitored for 12 weeks to study
their carrier status. In horses after the end of 3rd week all horses < 2 years of age
were found positive but at the end of 4th to 7th weeks there remained 5, 2, 1 and zero
out of 10, whereas in mules after the end of 2nd week all mules < 2 years of age were
positive but at the end of 3rd to 6th weeks there remained 7, 3, 1 and zero mules out
of 10, respectively on the basis of culture. But when compared the results with PCR,
at the end of the 4th week all horse <2 years of age were positive, but at the end of 5th
to 10th weeks there remained 7, 5, 4, 2, 1 and zero horses out of 10, while in mules at
the end of the 5th week all mules < 2 years of age were positive, but at the end of 6th
to 10th weeks there remained 9, 7, 3, 2 and zero mules out of 10, respectively. The
carrier status in 2 and 5 year old horses and mules was also evaluated. All horses
were found positive up to the 1st week but at the end of 2nd to 8th weeks there
remained 9, 7, 6, 3, 1, 1 and zero out of 10, whereas in mules all were positive up to
the 2nd week but at the end of 3rd to 7th weeks there were 6, 4, 2, 1, 1 and zero out of
10 mules, respectively on the basis of culture. But through PCR, all horses were
found positive up to 4th week but at the end of 5th to 9th weeks there were 9, 7, 6, 3,
2 and zero, in comparison with horses all mules were positive up to 5th week but at
the end of 6th to 10th weeks there were 8, 5, 2, 1 and zero. Horses and mules were
declared free of infection on the basis of three consecutive negative samples through
culture and PCR. Our findings are broadly consistent with the findings of Timoney,
1988 who reported that a 6 week course of shedding may be more typical. The
organism survives only for a short period in the environment unless protected in
moist discharges. Our study also correlates with the study of Timoney and Artiushin,
135
1997 who reported that the sensitivity of PCR appears to be much greater than the
culture. Our findings are in agreement with the results of Kahn, 2005 who reported
that most horses continue to shed organism up to one month following recovery.
Three negative nasopharyngeal swabs, at 4-7 days intervals, should be obtained prior
to release from quarantine, and a minimum isolation period should be one month.
Prolonged bacterial shedding has been identified in a small number of horses. Our
results were also broadly consistent with the findings of Sweeny et al., 1989 who
reported that the shedder state for S. equi implies that the equine harbors the S. equi
organism without manifesting overt clinical signs of strangles. Georage et al., (1983)
reported that 3 of 20 mares with strangles shed S. equi organism for at least 6 weeks
after lymph node rupture and a fourth mare never had lymphadenopathy, but on
arrival to the herd, it had cultures that were positive for S. equi and it continued to
shed S. equi intermittently over the next 10 months. Before this the longest reported
time between the disappearance of clinical signs of strangles and a culture positive
specimen of S. equi was 4 months. Our study correlates with the finding of Sweeny,
1990 who reported that horses with strangles may shed the organism for several
weeks following clinical recovery, with one survey detecting the organism for up to
10 month after exposure. Similarly Woolcock, 1975 suggested that the clinical
disease within a population might be a pre-requisite for development of the shedder
state because he was unable to isolate S. equi from horses on farms with strangles but
without active cases at the time of study. Sweeny et al., 1989 also reported failure to
isolate S. equi from horses which never developed strangles. It suggests that shedders
of S. equi among horses that never manifest clinical signs of strangles are rare. We
136
believe that S. equi shed in nasal secretions in horses recently recovered from
strangles is the most likely source of the organism for susceptible horses. Newton et
al., (2000) identified that after the clinical signs are abolished from the animals, the
animal remains in the carrier state and the most predominant site for S. equi was the
guttural pouch. He also reported that the prolonged carrier of S. equi, which lasted
upto 8 months, was again symmetrical with his study. Our results were also broadly
consistent with the findings of Sweeny et al., 2005 who reported that healthy horses
recovering from recent strangles disease might continue to harbor the S equi after a
full clinical recovery. There is evidence that a moderate proportion of horses continue
to harbor S equi for several weeks after clinical signs have disappeared, even though
the organism is no longer detectable in the majority 4 to 6 weeks after total recovery.
A recovered horse may be a potential source of infection for at least 6 weeks after its
clinical signs of strangles have resolved. Our results were also in line with the results
of Georage et al., (1983) who recorded that infected horses can shed S equi at least 4
weeks after the onset of clinical signs and the premises might harbor the organism for
a period of one year or longer. Although outbreaks may be initiated by the
introduction of clinically normal animals into a herd, diseases may become enzootic
on premises resulting in periodic outbreaks when the number of susceptible animals
increases.
It is concluded that do not mix recovered animals from strangles with healthy
animals at least for 9 weeks because the recovered animals remain carriers for
prolonged period of time (6-9 weeks). Periodic shedding of S equi can be a source of
infection for susceptible animals.
137
HAEMATOLOGICAL STUDIES:
The results of present study has revealed a significant increase (P < .05) of
total WBCs, MSNC, and basophils in strangles affected horses, while a non
significant difference was observed (P > .05) among values of lymphocytes,
eosinophils, basophils, erythrocytes, hemoglobin and packed cell volume. Whereas in
strangles affected mules the total WBCs, MSNC, and monocytes were significantly
increased (P < .05), while the values of lymphocytes significantly decreased (P < .05)
.Hemoglobin and Packed cell volume decreased but difference was non significant (P
> .05). A non significant difference (P>0.05) was observed in eosinophils, basophils
and erythrocytes. Our results correlate with the findings of Timoney, 2010 who
reported that hematological evidence of an acute phase response in the earlier stages
of strangles includes elevations in the white cell count (20750 ± 1583 cells/µl),
neutrophils (15058 ± 1604 cells/µl) and monocytes (6200 ± 1600 cells/µl) while
hemoglobin and pack cell volume may also be slightly or moderately reduced during
recovery. Gomez, 1990 also recorded hematlogic changes which include leukocytosis
with counts up to 30,000/µL, a segmented neutophil count that may be in excess of
25,000/µL. Hamlen et al., 1994 reported that strangles cases experienced leukocytosis
and neutrophilia in association with an increase in the health index but neutrophilia
was not associated with a left shift. Leukocytosis and neutophilia were also observed
in the foal experimentally inoculated with S .equi and have been reported by others
following experimental S. equi infections (Evers, 1968; Knight et al., 1975; Nara et
al., 1983). Our results were also in line with the findings of Higgins and Snyder
(2006) who reported pronounced leukocytosis with neutrophilia. Our results were
138
also broadly consistent with the findings of Hamlen et al., 1994 who reported that the
total leukocyte count increases quickly in horses in the first week post infection,
whereas this increase was delayed into the 2nd week. Other blood parameters
including total eosinophilic count, total basophilic count, total erythrocyte count,
packed cell volume, and hemoglobin concentration remained non significant (P > .05)
during the four weeks post infection. The results of this study were also in line with
Mahaffey, 1962 and Hamlen et al., 1994 who reported that packed cell volume in
mules did not differ in all the weeks. However it was less than the normal mules, as in
the present study. Higgins and Snyder, 2006 also reported leukocytosis and
neutrophilia. Our findings were also in the same line with the finding of Hamlen et
al., 1994 who reported that strangles cases had lower mean PCV and hemoglobin
concentration than non infected animals during weeks 4, 6 and 10 of the outbreak.
The erythrocytes count was also lower in +ve cases compared to non infected animals
during weeks 6 and 10. Collins, 1999 reported that a complete blood count proved to
be a useful adjunctive test to support a diagnosis of S equi and may help differentiate
horses with acute S equi infection from those with acute viral infection. Our findings
are also supported by the results of Evers, 1968 and Knight et al., 1975 who observed
decreases in PCV, hemoglobin concentration and erythrocytes counts have been
reported in horses following experimental inoculation with S. equi. Evers, 1968 also
observed that mean PCV, hemoglobin concentration and RBC count of 20 horses
decreased to 15%, 18% and 22%, respectively, 14 days post S. equi inoculation.
Knight et al., 1975 also reported that mean PCV and hemoglobin concentration
decreased to14% and 12%, respectively in 20 horses 6 days post infection. Evers,
139
1968 and Knight et al., 1975 both attributed these changes to in vivo S. equi
associated hemolysis. The most probable mechanisms responsible for the observed
mild to moderate decreases in PCV, hemoglobin concentration and erythrocytes count
in naturally exposed strangles and a more pronounced decrease seen in the
experimentally infected foals include factors affecting erythrocyte survival,
production and dilution. Hemodilution or increases in plasma volume, may account
for decreases in hematologic parameters of foals, particularly in the neonate and at
weaning (Harvey et al. 1987; Harvey et al. 1984; Jeffcott et al. 1982). Golland et al.,
1995 reported that hematological data was recorded for 8 horses. Leukocytosis with a
marked neutrophilia was present in 7 horses, with total nucleated cell counts of up to
23.2×109 cells/L. This was typically followed by the appearance of band neutrophils
and marked lymphocytosis. Seven horses were anemic (mean packed cell volume
0.32L/L). Evers, 1968 reported that temporary reduction of numbers of erythrocytes
and the amount of hemoglobin during the course of disease supports the theory that S.
equi exerts a hemolytic effect. Our findings are in agreement with the results of
Canfield et al., 2000 who reported that all experimentally infected horses with S. equi
showed a consistent leukocytosis as a consequence of a mature neutrophilia. These
changes developed within two days of infection and in some individuals persisted up
to 35 days. Neutrophilia in all the horses were mature and an increase in band
neutrophils was not detected in any of the horses. Other abnormalities in individual
horses included a mild lymphocytosis and a monocytosis.
140
BIOCHEMICAL STUDIES:
The result of the present study revealed that values of total serum protein,
serum globulin, and fibrinogen were significantly increased (P<0.05), whereas the
value of serum albumin significantly decreased (P<0.05) in strangles affected horses
and mules. Our study correlates with the study of Radostits et al. (2000) who reported
that hyperfibrinogenaemia is characteristic of both acute and chronic phase of disease.
Similarly hyperproteinemia attributable to a polyclonal gammaglobulinaemia is
characteristic of chronic abscess. Our studies also correlate with the findings of
Timoney, 2010 who reported that plasma protein (7.08±0.17g/dl and fibrinogen
(560±48mg/dL) are also increased. Fibrinogenaemia has also been reported by
Higgins and Snyder (2006). Gomez, 1990 also recorded fibrinogen level of 6.0mg/dL.
Our findings of fibrinogen correlate with the finding of Golland et al., 1995 who
recorded hyperfibrinogenaemia was present in 2 horses, with values of 7 and 8 g/L.
The results of the present study are in complete agreement with the findings of Taylor
and Wilson, 2006 who reported elevated concentration of globulin and fibrinogen,
and anemia of chronic inflammation are typical findings. Collins, 1999 reported that
plasma fibrinogen concentration frequently prove to be a useful adjunctive test to
support a diagnosis of S equi and may help differentiate horses with acute S equi
infection from those with acute viral infection.
THERAPEUTIC EFFICACY:
The resulst of in-vitro antibiotic sensitivity test revealed, that S equi was
sensitive to Procaine penicillin followed by ceftiofur Na, cephradine, erythromycin,
ampicillin, tetracycline, chloramphenicol, sulfamethoxazole, trimethoprim +
141
sulfanomides and gentamycin in equines whereas the results of in-vivo antibiotic
trials revealed that horses and mules suffering from strangles without abscess
formation were sensitive to Procaine penicillin followed by ceftiofur Na, cephradine
and erythromycin whereas those animals who developed abscess were ineffective.
From the results of of present study it is concluded that Procaine penicillin is most
effective in-vitro and in-vivo antibiotic followed by ceftiofur Na and cephradine.
These antibiotics might be used for the treatment of strangles infection. Our resulst
correlate with the study of George et al., 1983 who examined the sensitivity of in-
vitro antibiotics against S equi and found that all isolates were sensitive to following
antibiotics, penicillin, methicillin, erythromycin, tetracycline and ampicillin but
resistant to streptomycin and kanamycin. Our findings are also in agreement with the
results of Radostits et al. 2000 who recommended penicillin as a drug of choice for
treatment of strangles. Similarly, results of the present study are in agreement with
the findings of Higgins and Snyder, 2006 who reported that streptococci are very
sensitive to penicillin, ampicillin, erythromycin, chloramphenicol, cephalosporin and
tetracycline. Our results were also broadly consistent with the findings of Timoney,
2009 who reported that S. equi is highly sensitive to a wide range of antibiotics,
including Procaine penicillin, and there is no evidence of emerging drug resistance.
Penicillin treatment in the early part of the acute phase is often curative. Similarly,
results of the present study are in agreement with the findings of Manzoor et al., 2008
who reported antibiotic susceptibility of each of three Streptococcal species revealed
that in vitro Penicillin G and cefotoxime were very effective against Streptococci.
McAllister, 1982 and Swerezek, 1979 reported that antibiotic therapy is most
142
beneficial prior to the development of abscessation. McAllister, 1982 also observed
that penicillin is the drug of choice as there are no documented cases of S equi
resistance which is well in line with the results of present study. Our results were also
in agreement with the findings of Timoney, 1993 who reported that S. equi is very
sensitive to penicillin, chloramphenicol, erythromycin, tetracycline and lincomycin.
Procaine penicillin G is the antibiotic of choice and will show a quick clinical
improvement with reduction in fever and lymph node enlargement.
DISINFECTANT TRIALS:
Data regarding the comparison of different disinfectants is shown in table
4.53. Among four disinfectants, povidone iodine was found to be the best because its
phenol coefficient is 1.25 that is greater than phenol i.e. 1.00 while 0.6%solution of
H2SO4 showed a similar phenol coefficient as that of phenol. The phenol coefficients
of dettol and bleach were 0.5 and 0.75 respectively which are less than phenol i.e.
1.00. Therefore it is recommended that S. equi is highly sensitive to povidone iodine
and 0.6% H2SO4. Our results were also broadly consistent with the findings of
Higgins and Snyder (2006) who reported that all containers used for feed or water
should be cleaned and disinfected. Surfaces of stalls contaminated with discharges
should be similarly cleaned and disinfected. Effective disinfectants are povidone
iodine, chlorhexidine gluconate, 0.6 sulfuric acid, glutaraldehyde and phenol (1:200).
Our study also correlates with the study of Kahn, 2005 who used chlorhexidine
gluconate or gluteraldehyde for cleaning of contaminated equipment with S. equi.
Taylor and Wilson, 2006 reported that attempts should be made to disinfect areas that
have been occupied by infected horses. Organic material, such as feed and manure,
143
should be removed from the contaminated area and should be disposed of or
composted in an area that is not used for horses. Stall walls, feeders, waterers and
impervious floors should then be washed thoroughly to remove as much organic
material as possible before applying an effective disinfectant (eg, povidone iodine,
chlorhexidine and glutaraldehyde). Our results were also broadly consistent with the
findings of Jorm, 1992 who reported that organism is quickly killed at 560C and is
killed within 90 minutes by 0.6% sulfuric acid, a 1:200 dilution of phenol and by
disinfectants such as povidone iodine, chlorhexidine gluconate and glutaraldehyde.
Other disinfectants are less effective. Results of the present study are in agreement
with the findings of Wilson, 1988 and Reed et al., 2004 who reported that organism
does not survive for a prolonged period in the environment because of its
susceptibility to heat, sunlight, desiccation and many disinfectants including povidone
iodine, chlorhexidine gluconate and glutaraldehyde.
144
Chapter-06
SUMMARY
Strangles is an infectious malady of equidae characterized by upper
respiratory tract infection, dysponea, anorexia, regional suppurative lymphadenitis
and causes high morbidity and low mortality. Considering the significance and
utilization of equines in our country and the substantial losses rendered by Strangles,
the present project was designed to study epidemiology, diagnosis and chemotherapy
of strangles in Lahore and Sargodha districts of the Punjab province in Pakistan.
The present study comprised of five phases. In phase-I, epidemiology of the
disease including prevalence, variations in SeM, SzPSe and Se18.9 proteins and
mortality rate were studied in Lahore and Sargodha districts. For epidemiology, nasal
swabs and pus samples from the affected lymph nodes of 500 equines (n=250 horses,
n=250 mules) suspected for strangles were collected and cultured for identification of
S. equi. The collected samples were processed at Medicine and Microbiology
Laboratories of the University of Veterinary and Animal Sciences, Lahore, Pakistan
and Gluck equine research center, Department of Veterinary Science, University of
Kentucky, USA. Out of 250 horses and 250 mules, 113(45.2%) horses and 99
(39.6%) mules tested positive for S. equi. on the basis of culture. Number of S. equi
isolates were significantly higher (P<0.05) in pus samples taken from sub-mandibular
lymph nodes as compared to nasal discharge samples. The difference was significant
(P<0.05) among mules of different age groups. The highest prevalence of strangles
145
was recorded in horses and mules less than 2 year of age as compared to those having
age more than 2 years.
In the present study, prevalence of strangles round the year in horses and
mules were also calculated and it was found to be the highest during the months of
February, March, April and May while few cases were seen during the months of
January, June and July and no cases were observed during others months. The
significant difference was observed (p<0.05) among the prevalence levels of strangles
in different months of the year. Similarly when compared the prevalence of strangles
in different seasons of Pakistan i.e. summer, winter, spring and autumn. The highest
prevalence rate was recorded during the spring season.
The prevalence on the basis of Polymerase chain reaction (PCR) of S. equi in
horses and mules was also recorded. Out of 250 horses and 250 mules tested,
122(48.8%) horses and 113(45.2%) mules were positive for S. equi. When compared
the prevalence rate on the basis of PCR and culture of nasal and pus samples from
affected submandibular lymph nodes it revealed that the sensitivity of Polymerase
chain reaction appears to be much greater than culture. The culture along with PCR is
the best diagnostic technique for S. equi as PCR test does not differentiate between
dead and live bacteria, hence a positive test may not correlate with active infection;
therefore, a positive culture may be necessary to confirm the diagnosis.
In this phase of epidemiological study of disease, effect of selective pressure
of allelic diversity in SeM of S. equi on immunoreactive proteins SzPSe and Se18.9
was also studied. The aim of this study was to determine whether variations in SeM
are accompanied by variations in the immunoreactive surface of exposed SzPSe and
146
secreted Se18.9. Sequences of genes of 25 S. equi alleles isolated from different
countries of the world over a period of 40 years were compared. Twenty different
SeM alleles were identified including 6 not included in the data base (http://
pubmlst.org/szooepidemicus). Amino acid variation was also detected distal to the N-
terminus of SeM. No variation was observed in SzPSe except for an Australian isolate
which showed a deletion of one PEPK repeat. The Se18.9 protein in all 25 isolates of
S. equi did not exhibit any variation. Interestingly, only 2 SNP loci were detected in
Se18.9 compared to 93 and 49 in SeM and SzPSe respectively. The greater frequency
of mutation in SzPSe compared to Se18.9 may be related to a high rate of
recombination of SzPSe and the inclusion of exogenous DNA sequence based on the
atypical GC percentage of its central hyper variable region.
In horses the mortality rate was recorded as 1.64% whereas the mortality rate
in mules having less than 5 years of age was found to be 0.88%. No significant
difference (P>0.05) in mortality rate among horses and mules of different age groups
affected with strangles was observed.
In phase-II of the present study, carrier status of the horses and mules were
studied. Out of 122 horses found positive to PCR, 20 horses (10<2 years and 10
between 2 and 5 years of age) were selected and monitored for 12 weeks. Their nasal
swab samples were used for identification of bacteria through culture and PCR on
weekly basis. Till the end of 3rd week all horses < 2 years of age remained positive
but at the end of 4th to 7th weeks there remained positive only 5, 2, 1 and zero horses
out of 10, respectively on the basis of culture whereas through PCR at the end of the
4th week all horse <2 years of age were found positive, but at the end of 5th to 10th
147
weeks there remained 7, 5, 4, 2, 1 and zero horses out of 10, respectively. While all
the horses aging between 2 to 5 year, were positive up to the 1st week but at the end
of 2nd to 8th week out of 10 there were 9, 7, 6, 3, 1, 1 and zero horses respectively
positive on the basis of culture but through PCR, all horses were positive till 4th week
but at the end of 5th to 9th week number was reduced to 9, 7, 6, 3, 2 and zero.
Similarly, out of 113 mules, 20 mules (10<2 year and 10 between 2 and 5 years of
old) were also monitored for 12 weeks to study their carrier status. After the end of
2nd week all mules < 2 years of age were positive but at the end of 3rd to 6th weeks
there remained 7, 3, 1 and zero mules out of 10, respectively on the basis of culture
but through PCR at the end of the 5th week all mules <2 years of age were positive,
but at the end of 6th to 10th weeks there remained 9, 7, 3, 2 and zero mules out of 10,
respectively. While in 2 and 5 year old mules, all were positive up to the 2nd week
but at the end of 3rd to 7th weeks there were 6, 4, 2, 1, 1 and zero mules out of 10,
respectively on the basis of culture but through PCR, all mules were positive up to 5th
week but at the end of 6th to 10th weeks there were 8, 5, 2, 1 and zero. Horses and
mules were declared free of infection on the basis of three consecutive negative
samples through culture and PCR.
From the result of present study, it may be concluded that sensitivity of
Polymerase Chain Reaction appears to be much greater than culture for study of
carrier status of equines. Moreover, recovered animals should be kept in quarantine
period at least upto 9th week because the recovered horses and mules remain carrier
for prolonged period of time and can act as source of infection for susceptible animals
through periodic shedding of S equi.
148
In phase-III of the present study, effect of strangles on various haematological
parameters (Hemoglobin, Erythrocyte sedimentation rate, Total erythrocyte count,
Total leukocyte count, Differential leukocyte count, Packed cell volume) were
determined. Present study revealed that, total WBCs, MSNC, and basophils were
significantly increased (P < .05) in strangles affected horses, while the difference of
the values of lymphocytes, eosinophils, basophils, erythrocytes, hemoglobin and
Packed cell volume were non significant (P > .05). It was further found that in mules
total WBCs, MSNC, and monocytes in strangles affected mules were significantly
increased (P < .05), while the values of lymphocytes significantly decreased (P < .05).
Values of Hemoglobin and Packed cell volume were found to be decreased but
difference was non significant (P > .05). The difference of values of other blood
parameters like eosinophils, basophils and erythrocytes was observed non significant
(P>0.05).
The effect of strangles on total serum proteins, serum albumin, serum globulin
and fibrinogen were also studied in this phase. In the present study, the levels of total
serum protein, serum globulin, and fibrinogen were found to be significantly
increased (P<0.05), whereas a significant decrease (P<0.05) was observed in the
value of serum albumin in strangles affected horses and mules.
Phase-IV encompassed in-vitro and in-vivo antibiotic trials. In-vitro
antibiotic sensitivity of S. equi was determined against procaine penicillin,
erythromycin, chloramphenicol, ampicillin, cephradine, ceftiofur Na, tetracycline,
sulfamethoxazole, gentamycin and trimethoprim + sulfdiazine through disc diffusion
method. Four top ranking antibiotics were then administered to four groups
149
(comprising 10 horses and10 mules) for in-vivo trials. Efficacy of the antibiotics was
assessed weekly on the basis of negative nasal swab culture. Results of in-vitro
antibiotic sensitivity revealed that in horses and mules, S equi was most sensitive to
Procaine penicillin followed by ceftiofur Na, cephradine, erythromycin, ampicillin,
tetracycline, chloramphenicol, sulfamethoxazole, trimethoprim + sulfdiazine and
gentamycin whereas the result of in-vivo antibiotic trials revealed that horses and
mules suffered from strangles without abscess formation were most sensitive to
Procaine penicillin followed by ceftiofur Na, cephradine and erythromycin whereas
animals which developed abscess showed no response. It is concluded from the result
of present study that Procaine penicillin is most effective in-vitro and in-vivo
antibiotic followed by ceftiofur Na and cephradine. These antibiotics might be used
for the treatment of strangles infection.
Phase-V, comprised over in-vitro trials of disinfectants. Efficacy of
disinfectants, like povidone iodine, 0.6% H2SO4, dettol and bleach was assessed.
Phenol Co-efficient Test was applied, to ascertain efficacy of these disinfectants, used
in, in-vitro trials. Among four disinfectants, povidone iodine was found to be the best
one with a phenol coefficient of 1.25 that is greater than phenol i.e. 1.00 while 0.6%
H2SO4 showed similar phenol coefficient as that of phenol. The phenol coefficient of
dettol and bleach were observed as 0.5 and 0.75 respectively. Therefore it is
recommended that S. equi is highly sensitive to povidone iodine and 0.6% H2SO4.
150
LITERATURE CITED
Aguileta, G., G. Refregier, R. Yockteng, E. Fournier and T. Giraud, 2009. Rapidly
evolving genes in pathogens: Methods for detecting positive selection
and examples among fungi, bacteria, viruses and protists. Infect. Gen.
Evol. 9: 656-670.
Al-Ghamdi, G. M., V. Kapur, T. R. Ames, J. F. Timoney, D. N. Love and M. A.
Mellencamp 2000. Use of repetitive sequence-based polymerase chain
reaction for molecular epidemiologic analysis of Streptococcus equi
subspecies equi. Am. J. Vet. Res; 61(6):699-705.
Anonymous (2008-2009). Pakistan Economic Survey, Planning and Development
Division, Govt. of Pakistan, Islamabad. pp.32.
Anzai, T., A. Nakanishi, R. Wada , T. Higuchi , S. Hagiwara , M. Takazawa , K.
Oobayashi and T. Inoue,1997. Isolation of Streptococcus equi subsp.
equi from thoroughbred horses in a racehorse-breeding area of Japan.
J. Vet. Med. Sci; 59(11):1031-3.
Anzai, T., Y. Kuwamoto, R. Wada, S. Sugita, T. Kakuda, S. Takai, T. Higuchi, and
J.F. Timoney, 2005. Variation in the N-terminal region of an M-like
protein of Streptococcus equi and evaluation of its potential as a tool in
epidemiologic studies. Am. J. Vet. Res. 66: 2167-71.
Ashraf, M. ,2000. Some Epidemiological and Clinicopathological Studies on
Strangles in Horses and Mules. PhD Thesis, University of Agriculture,
Faisalabad, Pakistan.
151
Bazeley, P. L., 1943. Studies with equine Streptococci 5: Some Relations between
virulence of Streptococcus equi and immune response in the host. Aus.
Vet. J; 19:62-85.
Bazeley, P. L. and J. Battle, 1940. Studies with equine Streptococci. I: A Survey of
Beta Haemolytic Streptocci in Eqine Infections. Aus. Vet. J; 16: 141-
146.
Boschwitz, J.S., J. F.Timoney, 1994. Inhibition of C3 deposition on Streptococcus
equi subsp. equi by M protein: a mechanism for survival in equine
blood. Infect. Immun. 62: 3515–3520.
Boyes, S. M. B., R. L. Young, D. D. Canton and F. C. Mohr, 1991. Streptococcus
equi infections as a cause of panophthalmitis in a horse. Equine Vet.
Sci; 11 (4): 229-231.
Bryans, J. T. and B. O. Moore, 1972. Group C streptococcal infections of the horse.
In: Wannamaker LW, Masten JM (ed): Streptococci and Streptococcal
Diseases: Recognition, understanding and Management. London,
Academic Press, 327-337.
Bryans, J. T., E. R. Doll and B. P. Shephard, 1964. The etiology of strangles.
Cornell Vet. 54: 198-205.
Buxton, A. and G. Fraser. 1975. Animal microbiology. Blackwell Scientific
Publication London: 165-177.
Canfield, P. J., D. N. Love, J. Rainger and G. D. Bailey, 2000. Strangles in horses. A
report for the rural industries research and development corporation.
RIRDC Publication No 00/7, Project No. US-24/A.
152
Cappuccino, J. G. and N. Sherman 2004. Microbiology a laboratory manual. 6th
edition: 280-281.
Castagnoli, B. and A. Balboni,1942. Cerebral localization of Streptococcus equi Riv.
Milit. Med. Vet. Roma; 5: 69-82.
Chanter, N., N. C. Talbot, J. R. Newton, D. Hewson, and K. Verheyen, 2000.
Streptococcus equi with truncated M-proteins isolated from outwardly
healthy horses. Microbiology 146:1361–1369.
Clabough, D., 1987. Streptococcus equi infection in the horse: a review of clinical
and immunological considerations. Equine Vet. Sci; 7:279-282.
Coles, E. H., 1980. Veterinary clinical pathology. 3rd ed. W. B. Saunders Company,
London: 40-70.
Collins, M. J. and U. Wernery, 1999. Control and eradication of a Streptococcus equi
outbreak on a horse riding establishment in the United Arab Emirates.
Equine infectious diseases VIII. Newmarket, R& W Publications
Limited.
Dalgleish, R., S. Love , H. M. Pirie , M. Pirie , D. J. Taylor and N. G. Wright , 1993.
An outbreak of strangles in young ponies. Vet. Rec; 132(21):528-31.
De Lahunta, A. 1977. Neurological problems in the horse. Proceedings of the 19th
Annual American Association of Equine Practitioners. pp. 25-32.
Dwyer, R. M., 1995. Disinfecting equine facilities. Rev. Sci. Tech; 14(2): 403-18.
Eberbeck, E. and A. H. Halswick, 1943. Pathogenesis of the sequelae of equine
infectious bronchitis and strangles. Arch. Wiss. Prakt. Tierheilk; 78:
334-351.
153
Ensink, J. M., J. A. Smit and E. V. Duijkeren, 2003. Clinical efficacy of
trimethoprim/sulfadiazine and procaine penicillin G in a Streptococcus
equi subsp. zooepidemicus infection model in ponies. J. Vet.
Pharmacol Ther; 26(4):247-52.
Evermann, J. F., A. C. Ward and J. V. Schalie, 1987. Equine influenza outbreak.
Journal of Equine Vet. Sci; 7 (1): 43-44.
Evers, W. D., 1968. Effect of furaltadone on strangles in horses. J. A. V. M. A; 152:
1394-1398.
Fallon, E. H., 1969. The clinical aspects of streptococcal infections of horses. J.A. V.
M. A; 155:413-414.
Fan, H., Y. Wang, F. Tang and C. Lu, 2008. Determination of the mimic epitope of
the M-like protein adhesin in swine. BMC Microbiol. 8: 170.
Ford, J. and M. D. Lokai, 1980. Complications of Streptococcus equi infection.
Equine Pract; 2: 41-44.
George, J. L., J. S. Reif , R. K. Shideler , C. J. Small , R. P. Ellis , S. P. Snyder and A.
E. McChesney , 1983. Identification of carriers of Streptococcus equi
in a naturally infected herd. J. Am. Vet. Med. Assoc; 183(1):80-4.
Golland, L. C., D. R. Hodgson., R. E. Davis., R. J. Rawlinson., M. B. Collins., S.A.
McClintock and D. R. Hutchins, 1995. Retropharyngeal lymph node
infection in horses: 46 cases (1977-1992). Aust. Vet. J; 72 (5): 161-
164.
154
Gomez, H. J., 1990. The epidemiologic and immunologic characterization
Streptococcus equi in foals (Thesis). Saskatoon, Saskatchewan,
University of Saskatchewan.
Gronbaek, L. M., O. Angen , H. Vigre and S. N. Olsen, 2006. Evaluation of a nested
PCR test and bacterial culture of swabs from the nasal passages and
from abscesses in relation to diagnosis of Streptococcus equi infection
(strangles). Equine Vet J; 38(1):59-63.
Gumbrell, R., 1987. Strangles in horses. Surveillance, New Zealand. 14 (4): 11-12.
Hamlen, H. J., J. F. Timoney and R. J. Bell, 1992. Hematologic parameters of foals
during a strangles epizootic. Equine Veterinary Science; 12:86-92.
Hamlen, H. J., J. F. Timoney and R. J. Bell, 1994. Epidemiologic and immunologic
characteristics of Streptococcus equi infection in foals. J. Am. Vet.
Med. Assoc; 204(5):768-75.
Harrington, D. J., I. C. Sutcliffe, and N. Chanter, 2002. The molecular basis of
Streptococcus equi infection and disease. Microbes and Infection; 4
(4): 501-510.
Harvey, J. W., R. L. Asquith, P. K. McNulty, J. Kivipelto and J. E. Bauer, 1982.
Hematology of foals up to one year of age. Equine Vet. J; 16: 347-353.
Harvey, J. W., R. L. Asquith, W. A. Sussman and J. Kivipelto, 1987. Serum ferritin,
serum iron, and erythrocyte values in foals. Am. J. Vet. Res; 48:1348-
1352.
Heather, Z., M.T.G., K.F. Holden, J. Steward, L. Song, G.L. Challis, C, Robinson, C.
Davis-Poynter and A.S.Waller, 2008. A novel streptococcal integrative
155
conjugative element involved in iron acquisition. Molecular
Microbiology. 70 (5): 1274 – 1292.
Higgins, A. J. and J. R. Snyder, 2006. The equine manual. W.B. Saunders Co.,
Philadelphia.2nd edi. pp 75-81.
Hignett, S. L. and W. S. King. 1940. Streptococcal infection in the commercial horse
.Vet. J; 96: 81-84.
Jannatabadi, A. A., G. R. Mohammadi, M. Rad and M. Maleki. 2008. Molecular
identification of Streptococcus equi subsp. Equi and Streptococcus
equi subsp. Zooepidemicus in nasal swabs samples from horses
suffering respiratory infections in Iran. Pak. J. Biol. Sci; 11: 468-471.
Jeffcott, L. B., P. D. Rossdale and D. P. Leadon, 1982. Hematologic changes in the
neonatal period of normal and induced premature foals. J. Repro. Fert;
32:537-544.
Jorm, L. R., 1990. Strangles in horse studs: incidence, risk factors and effect of
vaccination. Aust. Vet .J; 67(12):436-9.
Kahn, C. M., 2005. The merck veterinary manual. Merck & Co.,INC. Whitehouse
station, N. J. USA. 9th edi. Pp-1213.
Kelly, C., M. Bugg, C. Robinson, Z. Mitchell, N. Davis-Poynter, J.R. Newton, K.A.
Jolley, M.C Maiden, M.C. and A.S. Waller, 2006. Sequence variation
of the SeM gene of Streptococcus equi allows discrimination of the
source of strangles outbreaks. J. Clin. Microbiol. 44: 480–486.
156
Knight, A. P., J. L. Voss, A. E. McChesney and H. G. Bigbee. 1975. Experimentally-
induced S. equi infection in horses with resultant guttural pouch
empyema. VM/SAC Equine Pract; 1194-1199.
Knowles, E. J., T. S. Mair, N. Butcher , A. S. Waller and J. L. Wood, 2010. Use of a
novel serological test for exposure to Streptococcus equi subspecies
equi in hospitalised horses. Vet. Rec; 166(10):294-7.
Kumar, P., Muthupalani, S., Timoney, J.F. 2007. Unpublished Data.
Mahaffey, L.W., 1962. Respiratory conditions in horses. Vet. Rec; 74 (47): 1295-
1306
Manninger, R., 1949. Aetiology of stangles. Acta Vet. Hung; 1: 73-75.
Manzoor, S., M. Siddique, S. U. Rahman, and M. Ashraf, 2008). Occurance of
lancefield group C Streptococcal species in strangles cases of Folas in
Punjab, Pakistan. Pakistan vet. J. 28:17- 20.
Mayr, A., 1987. Respiratory infectious diseases in horses. Tierarztl-Prax-Suppl; 2: 1-
4.
McAllister, E. S., 1982. Bacterial respiratory infections, in: Mansmann RA,
McAllister ES (ed): Equine medicine and surgery, ed 3. Santa Barbara,
American Vet. Publication Inc; pp 734-737.
McGee, W. R., 1969. The clinical aspects of streptococcal infections of the horse, in:
Proceedings 2nd Int. Conf. Equine infectious Diseases, Paris; 227-230.
Merchant, I. A. and R. A. Packer, 1983. Veterinary bacteriology and virology. 7th ED.
CBS. Publisher London: 62-72 and 112.
157
Minett, F. C., 1944. The problem of equines strangles in India. II Climatic factors as
related to the incidence of equine strangles in India. Indian J. Vet. Sci;
14: 1-12 and 75-94.
Moore, B. O., Bryans, J. T., 1969. Antigenic classification of group C animal
streptococci. J.Am. Vet. Med. Assoc. 155, 46-21.
Muhktar, M. M. and J. F. Timoney,1988. Chemotactic response of equine
polymorphonuclear leucocytes to Streptococcus equi. Res. Vet. Sci; 45
(2): 225-229.
Namikawa, S., Y. Kurihawa and M. Emori, 1940. on the specificity of Streptococcus
equi from view point of Fortner’s Anaerobic Cultivation. Japan J.
Vet. Sci; 2: 518-530.
Nara, P. L., S. Krakowka, T. E. Powers and R. C. Garg, 1983. Experimentally S. equi
infection in the horses: Corelation with in vivo and in vitro immune
responses. Am. J. V. Res; 44:529-534.
Natarajan, D. and L. Ingeborg, 2003. Purdue University Animal Disease Diagnostic
Laboratory Newsletter. http://www.addl.purdue.edu/newsletters/
2003/Spring/strangles.shtml (2 November 2004).
Newton, J. R., Wood, J. L., Dunn, K. A., DeBrauwere, M. N. and Chanter, N,1997.
Naturally occurring persistent and asymptomatic infection of the
guttural pouches of horses with Streptococcus equi. Vet. Rec. 140: 84–
90.
Newton, J. R., K. Verheyen, N. C. Talbot , J. F. Timoney , J. L. Wood , K. H.
Lakhani and N. Chanter, 2000. Control of strangles outbreaks by
158
isolation of guttural pouch carriers identified using PCR and culture of
Streptococcus equi. Equine Vet J ; 32(6):515-26.
Niebauer, G. W., G. Punzet and R. Swoboda, 1979. Metastatic strangles abscess in
the brain of a horse. Wiener. Tierartztliche Monatsschrift; 66 (11):
321-325.
Paunovic, S., 1957. A malignant form of strangles. Vet. Glasn; 11: 617-619.
Peatt, E. S. W., 1945. The Army Veterinary Services in India and Burma. Vet. Rec;
57: 219-221.
Piche, C. A., 1984. Clinical Observations on an Outbreak of Strangles. Can. Vet. J;
25(1):7-11.
Prescott, J. F., S. K. Srivastava , R. deGannes and D. A. Barnum ,1982. A mild form
of strangles caused by an atypical Streptococcus equi. J. Am. Vet. Med.
Assoc; 180(3):293-294.
Quinn, P.J., M.E Carter, B. Markey, and G.R. Carter, 1994. The streptococci and
related cocci. In: Clinical Veteterinary Microbiology, Wolfe
publishing, Mosby-year Book Europe Limited, London. pp 127-136.
Radostits, O.M., C.C. Gay, D.C. Blood, and K.W. Hinchcliff, 2000. Veterinary
Medicine. A Text Book of the Diseases of Cattle, Sheep, Pigs, Goats
and Horses. W.B. Saunders Co., Philadelphia. pp 705.
Reed, S. M., W. M. Bayly and D. C. Sellon, 2004. Streptococcus equi infection
(strangles), in Equine Internal Medicine. St. Louis, MO, Saunders, pp
308-312.
159
Roberts, S. R., 1971. Chorioretinitis in a band of horses. J. Am. Vet. Med. Ass. 158:
2043-2046.
Shultz, J. W., 1888. The Streptococcus of strangles. The Journal of Comparative
Pathology and Therapeutics; 1:(3) 191-208.
Sonea, I. M., 1984 Strangles, in Merchant IA (Ed): Veterinary Bacteriology and
Virology, Ames, IA, Iowa State University Press, pp 590-592.
Srivastava, S. K. and D. A. Barnum, 1981. "The serological response of foals to
vaccination against strangles." Can J. Comp. Med; 45(1): 20-25.
Srivastava, S. K. and D. A. Barnum, 1983a. Vaccination of pony foals with M-like
protein of Streptococcus equi. American Journal of Veterinary
Research; 44 (1): 41-45.
Sweeney, C. R., C. E. Benson , R. H. Whitlock , D. A. Meirs , S. O. Barningham , S.
C. Whitehead and D. Cohen, 1989. Description of an epizootic and
persistence of Streptococcus equi infections in horses. J. Am. Vet.
Med. Assoc; 194(9):1281-6.
Sweeney, C. R., R. H. Whitlock and Meirs, 1987. Complications associated with
Streptococcus equi infection on a horse farm. J. Am. Vet. Med. Assoc;
191:1446-1447.
Sweeny, C. R., 1990. Streptococcus equi. In smith B, ed: Large animal internal
medicine. St Louis, Mosby-year book, pp, 519.
Swerezek, T. W., 1979. Aggravation of strangles, equine clostridial typhlocolitis
(colitis x) and bacterial venereal diseases in the horse by antibacterial
drugs. Proc. AAEP; 25:305-311.
160
Tajima, M. and A. Ueda, 1953. The microscopical observations on the four cases of
purulent encephalomyelitis in horses affected with strangles. Vet. Res.
Japan; 1: 41-48.
Taylor, S. D. and W. D. Wilson, 2006. Streptococcus equi subsp. equi (Strangles)
infection. Clinical Techniques in Equine Practice; 211-217.
Thrusfield, M., 2002. Veterinary epidemiology. 2nd Ed. Blackwell publisher London.
Timoney J. F., 2010. Strangles. In Equine Reproduction, 2nd Ed. A. McKinnon, E.
Squires, D.Varner,
Timoney, J. F., 1988. Protecting against stangles: A contemporary view. Equine Vet.
J; 20:392.
Timoney, J. F., 1988. Shedding and maintenance of Streptococcus equi in typical and
atypical strangles, in: Powell D. (Ed): Equine Infectious Disease V:
Proceedings of the Fifth International Conference. Kentucky, the
University of Kentucky Press, pp 28-33.
Timoney, J. F., 1993. Strangles. Vet. Clin. North. Am. Equine Pract; 9(2):365-74.
Timoney, J. F., 2004. The pathogenic equine streptococci. Vet. Res; 35(4):397-409.
Timoney, J. F., 2009. Current therapy in equine medicine. W.B. Saunders Co.,
Philadelphia. 6th edi. pp 128.
Timoney, J. F. and J. Trachman, 1985. Immunologically Reactive Proteins of
Streptococcus equi. Infection and immunity; 48:29-34.
Timoney, J. F. and S. C. Artiushin, 1997. Detection of Streptococcus equi in equine
nasal swabs and washes by DNA amplification. Vet Rec; 141:446-447.
161
Timoney, J. F., S. C. Artiushin and J. S. Boschwitz, 1997. Comparison of the
Sequences and Functions of Streptococcus equi M-Like Proteins SeM
and SzPSe. Infection and immunity; 65: 3600–3605.
Timoney, J.F, R. DeNegri, A. Sheoran, N. Forster, 2009. Affects of N-terminal
variation in the SeM protein of Streptococcus equi on antibody and
fibrinogen binding. Vaccine. 28:1522–1527.
Timoney, J.F, A. Qin, S. Muthupalani, S. C. Artiushin, 2007. Vaccine potential of
novel surface exposed and secreted proteins of Streptococcus equi.
Vaccine. 25:5583–5590.
Tiwari, R., A. Qin, S.C. Artiushin, J.F.Timoney,2007. Se18.9, an anti-phagocytic
factor H binding protein of Streptococcus equi. Vet. Microbiol. 121:
105–115.
Todd, A. G., 1910. Strangles. Journal of Comparative Pathological and
Therapeutics; 23:212- 229.
Tuji, Y. and A. Sato, 1940. On the type Differentiation of Hemolytic Streptococci
from Strangles, Pneumonia and other Streptococcal Diseases of
Equines. The type Differentiation by Biological examination. Japan
J. Vet. Sci; 2: 593-616.
Valentine-Weigand, P., G. S. Chhatwal and H. Blobel, 1988. Adherence of
streptococcal isolates from cattle and horses to their respective host
epithelial cells. American Journal of Veterinary Research; 9: 1485-
1488.
162
Van Dorssen, 1939. Aetiology of Benign Strangles. Tijdschr. Diergeneesk. 66:
716-730.
Verheyen, K., J. R. Newton , N. C. Talbot , M. N. de. Brauwere and N. Chanter,
2000. Elimination of guttural pouch infection and inflammation in
asymptomatic carriers of Streptococcus equi. Equine Vet. J; 32(6):527-
32.
Vukovic, V., 1961. Strangles in Sarajevo during 1952-1959. Veterinaria, Sarajevo;
10: 125-128.
Wagenaar, G. and A. Van. Der. Schaaf. 1965. Strangles. Tijdschr. Diergeneesk; 90:
315-323.
Walker, J.A. and J.F. Timoney, 1998. Molecular basis of variation in protective SzP
proteins of Streptococcus zooepidemicus. Am. J. Vet. Res. 59: 1129-
33.
Waller, A. S. and K. A. Jolley, 2007. Getting a grip on strangles: recent progress
towards improved diagnostics and vaccines. Vet. J; 173(3):475-6.
Weichselbaum, T.E., 1946. An accurate and rapid method for the determination of
proteins in small amounts . Am. J. Clin. Pathol. Tech. Suppl., 10: 40-
49.
Wilson, W. D., 1988. Streptococcus equi infections (strangles) in horses. Equine
Pract; 10:12-18.
Wisecup, W. G., C. Schroder and N. P. Page ,1967. Isolation of Streptococcus equi
from burros. J. Am. Vet. Med. Ass; 150: 303-306.
163
Wood, J. L., K. Dunn, N. Chanter and N. de Brauwere, 1993. Persistent infection with
Streptococcus equi and the epidemiology of strangles. Vet. Rec. 133:
375–375.
Woolcock, J. B., 1975. Epidemiology of equine streptococci. Res Vet Sci; 18(1):113-
4.
Woolcock, J. B., 1975. Studies in atypical Streptococcus equi. Res. Vet. Sci. 19(2):
115-9.
Yelle, M. T., 1987. Clinical aspects of Streptococcus equi infection. Equine Vet. J; 19
(2): 158-162.
Zadeh, F. N., F. G. A. Pour and S. M. Khajeh-Nasiri, 1992. Epizootological
investigation of strangles in the equine stables in Tehran. J. Equine
Vet. Sci; 12(6): 401-402.