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Original Article
Five-year update on the mouse model of orthotopic lung transplantation:
Scientific uses, tricks of the trade, and tips for success
1Department of Surgery, 2The First Affiliated Hospital of Sun Yat-sen University, Guangzhou 510080, China; 3Center of Chest Medicine and Surgery, Ehime University, Ehime, Japan; 4Department of Thoracic Surgery, Okayama University Hospital, Okayama, Japan; 5Department of Pathology and Immunology, Washington University School of Medicine, USA
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Abstract
It has been 5 years since our team reported the first successful model of orthotopic single lung transplantation in the mouse (1).
There has been great demand for this technique due to the obvious experimental advantages the mouse offers over other
large and small animal models of lung transplantation. These include the availability of mouse-specific reagents as well as
knockout and transgenic technology. Our laboratory has utilized this mouse model to study both immunological and nonimmunological
mechanisms of lung transplant physiology while others have focused on models of chronic rejection (1-19).
It is surprising that despite our initial publication in 2007 only few other laboratories have published data using this model
(20-25). This is likely due to the technical complexity of the surgical technique and perioperative complications, which can
limit recipient survival (9,26). As two of the authors (XL and WL) have a combined experience of over 2500 left and right
single lung transplants, this review will summarize their experience and delineate tips and tricks necessary for successful
transplantation. We will also describe technical advances made since the original description of the model (1,5,6).
Key words
Mouse model; orthotopic lung transplantation; surgical complications; evaluation
J Thorac Dis 2012;4(3):247-258. DOI: 10.3978/j.issn.2072-1439.2012.06.02 | ||||||||||||||||||||||||||||||||||||
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Introduction
As of 2011 our laboratory has successfully completed over 4,500
mouse orthotopic lung transplants as a part of investigatorinitiated
studies and collaborations with biotechnological
companies (1-7,27-30). While in the beginning of our experience
technical obstacles were encountered, after an initial learning
curve complications are now rare (less than 1%, not including
post-transplantation atelectasis). Other laboratories have
reported technical success rates of around 80-90% (9,26). As we
have already described the rudimentary steps of this procedure
(1,5) , here we will focus only on major surgical complications
that hinder experimental success. We will also discuss the causes
of technical failures and steps that can be taken to prevent them. | ||||||||||||||||||||||||||||||||||||
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Complications
Pneumothorax
The first type of pneumothorax (Type I pulmonary injury)
stems from technical errors that occur due to excessive handling
of the lung. This results in a severe air leak, mostly because of
a bronchial tear, or damage to the lung surface. This becomes
evident immediately after graft reperfusion and reaeration. This
type of injury is worth repairing if the area of injury is small or
the recipient is a very rare transgenic mouse. Although the mouse
bronchus and lung tissue are very friable, one can still repair this
form of damage by placing a shorter cuff on the torn bronchus to
incorporate the area of damage into the cuff or utilize 10-0 nylon
suture (Sharpoint AK-D106, Surgical Specialties Corporation,
Figure 1) to repair a small hole in the donor or recipient
bronchus. As for tiny defects on the graft surface one can use
tissue glue to seal it (Webglue, Webster Veterinary Supply Inc,
Figure 2). In our experience a successful repair of this type of
injury is easy to achieve and the graft can often be salvaged.
Figure 1. 10-0 Nylon for suturing tiny hole of bronchus.
Figure 2. Animal Surgical Glue for sealing small air leak of graft lung.
The second type of pneumothorax (Type II pulmonary injury)
is more dangerous because the lung injury is small and may not
be evident immediately upon release of the hilar structures.
This is usually discovered only after extubation of the animal
and is often manifested by respiratory distress and difficulty
in initiating spontaneous respirations. Without immediate
reintubation the recipient will die within 1 or 2 minutes and
is unlikely to be resuscitated. In order to salvage this situation
the animal must be quickly reintubated and the upper part of
the abdomen opened. This will allow immediate diagnosis of
the problem as either the left or the right side of the diaphragm
will bulge down with an obvious tension pneumothorax. In
our experience a Type II pulmonary injury can occur either on
the side of the transplant or native lung. While the reason for
this injury can be difficult to determine it may be related to the
process of mechanical ventilation. In the mouse the respiratory
rate varies between 60-230 breaths per minute (bpm). The
tidal volume can vary between 0.09-0.38 mL/breath resulting
in a minute ventilation of 11-36 mL/min (31). Despite this
physiology our preferred mouse ventilator settings include a
tidal volume of 0.5 mL and a respiratory rate of 110-120 bpm,
which yield minute ventilations exceeding the normal range.
Other laboratories performing similar procedures recommend
even higher tidal volumes (26). In our experience decreasing
the minute ventilation for extended portions of the procedure
leads to poor long-term outcome (unpublished data). During
single lung ventilation, however, this high volume positive-pressure
may cause barotrauma and volutrauma, which may lead to a Type
II pulmonary injury (32). While serious, this complication is
very rare (from our database we have encountered fewer than
20 cases in 5 years), and the severity of pulmonary damage
is unpredictable. In order to prevent this complication we: (I)
reduce the tidal volume from 0.5 mL to 0.35-0.40 mL when
single lung ventilation is initiated and maintain this low tidal
volume until the lung graft is implanted and (II) inflate the lung
graft for no more than 1-2 seconds after implantation. However,
if a pneumothorax does occur the only possible treatment is to
reintubate the animal immediately, incise the upper abdomen
and cut a small window in the diaphragm to release the air
that has collected under tension in the chest. After recipient
hemodynamics have stabilized the diaphragmatic window is left
open and the abdominal incision is closed (Figure 3). If needed,
one can close the diaphragm window at a later time point.
However, it is unlikely that these salvage maneuvers will result in
a ventilated lung at the time of sacrifice as atelectasis will occur
after this type of injury. Therefore, mice can generally not be
used for data generation if this complication arises.
Figure 3. Diaphragm window for treatment of type II pulmonary injury.
Graft atelectasis
The anatomy of the mouse lung differs from that of other species
(33). Total lung capacity (TLC) of the mouse is about 1ml
compared to 10 ml in the rat. The alveoli of the mouse lung are
significantly smaller than those of the rat (34) and the airways
constitute a large percentage of the lung volume in the mouse
(11%) compared to the rat (5.7%). Cartilage is present in the
mouse trachea, but is less well organized than in other species.
Mouse lungs also have fewer respiratory bronchioles and airway
generations than humans. Two other significant features of
the mouse lung anatomy are the thinness of the respiratory
epithelium and the relatively large airway lumen (35). This
large airway caliber may reduce the flow-resistive load in this
small animal with a rapid respiratory rate (33). All of these
characteristics work in favor of preventing atelectasis of the
mouse lung in comparison to other animal models. Nevertheless,
it has been brought to our attention that non-alloimmunemediated
collapse of the grafted lung, which is not associated
with graft rejection, but rather due to technical problems, is the
most common complication encountered by other laboratories.
Thus, in our opinion, pulmonary atelectasis after transplantation
is generally due to methodologic problems rather than inherent
anatomic considerations.
Based on our experience and serial sacrifice, pulmonary
collapse that is unrelated to graft rejection usually occurs on
the third to fifth post-operative day after transplantation and is
often not diagnosed until the time of sacrifice. The diagnosis of
atalectasis is further complicated by the fact that mice do not
manifest any outward signs of shortness of breath when this
occurs (6). Furthermore, while non-invasive diagnostic tools
such as small animal MRI are available to monitor pulmonary
graft function after transplantation. the routine use of such
modalities is impractical (Figure 4) (29). Thus, we contend
that the best method for preventing graft atelectasis is to pay
meticulous attention to surgical details and rely on several “tricks”
that we have accumulated over the years in order to prevent this
complication.
Figure 4. MRI images obtained 3 days after lung transplantation reveal ventilation of lung graft on three serial coronal images (top) and graft
atelectasis in a different animal (bottom). As previously described (29) aerated lung looks black on MRI due to lack of signal intensity while
a consolidated lung looks white due to the high intensity of this water-rich structure when atelectatic.
In our mouse model, we believe pulmonary collapse occurs
due to two main reasons. The first one involves pathophysiologic
changes in pulmonary parenchyma after transplantation and
the second one is technical failure. Graft ischemia-reperfusion
injury alters graft compliance, increases air flow resistance and
contributes to atelectatic collapse of the lung (36). Anastomotic
problems of the small mouse bronchus also can lead to
atelectasis and anastomotic ischemia followed by remodeling
and granulation tissue formation, which leads to bronchial wall
thickening and significant airway obstruction (37-39). Technical
problems due to poor orientation of the cuff, donor to recipient
cuff mismatch and other mishaps can lead to atelectatic collapse
of the lung graft as well. While some have demonstrated that
cuffing the bronchial anastomosis can be problematic and
may lead to airway stenosis in the rat (36) the small size of the
bronchial structures in the mouse makes suturing this structure
nearly impossible (unpublished observations). Thus, while
retaining the cuff technique for the bronchial anastomosis we
have gradually modified the surgical approach and have made
significant revisions in the method for cuffing this structure.
First, we now shorten the donor bronchus as close as possible
to the secondary carina in order to reduce anastomotic ischemia
and improve the bronchial blood supply (40-43). This type of
maneuver also results in the need to shorten the venous cuff
in order to prevent rotation and torsion of the hilar structures.
Second, we use as big a cuff as possible in order to reduce air
flow resistance (with gain of experience some microsurgeons
now cuff most strains of mice with a cuff made from a #18
angiocatheter instead of a #20 as we had previously described
(5) (Figure 5). As the hilar anatomy of each strain differs (C3H
and CBA/Ca mice for example have comparably foreshortened
hilar structures) the size of the cuff must match each bronchus
in order to achieve not only optimal ventilation, but also prevent
venous complications (see below). After these modifications our
incidence of graft atelectasis has decreased to <5%.
Figure 5. Adjustment of bronchus cuff size. From L to R, the size
is 18#, 20# and 24#, which are used for bronchus, pulmonary vein,
and pulmonary artery, respectively.
Pulmonary venous thrombosis
This complication does not usually lead to immediate postoperative
demise and is usually discovered at autopsy either
due to recipient death before the scheduled sacrifice or at the
time of sacrifice. Gross appearance of the graft can be diagnostic
of pulmonary venous thrombosis as the lung is dark red or
black, is grossly enlarged and firm. Often a clot can be seen in
the pulmonary vein. At times the right upper lobe of the native
lung can be involved as well due to propagation of clot. Venous
thrombosis is usually caused by pulmonary vein torsion or
compression of the venous outflow from a large bronchus cuff.
Thus, one must balance the “trade off” of a large bronchial cuff, which results in a lower incidence of graft atelectasis, but a higher
incidence of pulmonary venous thrombosis. Nevertheless, the
incidence of this complication can be reduced to very low levels
by meticulous attention to detail in preventing venous torsion
and proper cuffing of the hilar structures to reduce rotation of
the lung by an imbalance between the bronchial and venous
cuffs. | ||||||||||||||||||||||||||||||||||||
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Evaluation
Evaluation of technical success and rejection
After technically successful lung transplantation the graft
should be well aerated and perfused, even in the face of mild
acute rejection. As mentioned above, unlike cardiac allografts,
where early technical failure can be manifested by cessation
of heart beat, the evaluation of the technical success of lung
transplantation requires either gross inspection after sacrifice or
radiographic imaging. Conventional imaging modalities, such
as X-rays, lung perfusion scintigraphy or angiography may be useful in humans and large animals, but are not practical for large
throughput experiments in mice (44-48). Either the resolution
of these techniques is not suitable to evaluate subtle pathological
changes, or the techniques cannot be performed on a serial
basis in the same animal (36). Greschus suggested Flat-Panel
Volumetric Computer Tomography (fpVCT) as a precise tool
to assess the success of rat orthotopic lung transplantation that
can be used to follow the process of graft rejection with very high
spatial resolution (36). Such a technique, however, might not be
applicable to the mouse due to its small size. We have recently
focused on small animal MRI to evaluate pulmonary pathology
and have found this technique to be a highly reproducible
non-invasive approach to visualize anatomic pathology of the
lung such as pulmonary collapse due to either advanced acute
rejection or atelectasis (29) (Figure 4). However, we realize that
this modality might not be easily accessible to all laboratories
and thus to date there still is no easily accessible, effective
and precise imaging modality to evaluate the grafted lung for
technical success or rejection. Our current practice thus relies on
grading rejection based on strict histologic criteria, which mirror
human graft evaluation. In 1990, the International Society for
Heart and Lung Transplantation (ISHLT) adopted a “Working
Formulation of the Standardization of the Nomenclature in the
Diagnosis of Lung Rejection,” which was revised in 1996 and
again in 2007 (49). We have thus adapted this grading scale for
the mouse model (Table 1). The development of better imaging
modalities in the future may facilitate serial monitoring of graft
outcome and early detection of technical failures.
Evaluation of graft function
Mouse orthotopic lung transplantation is a very useful model
not only because it mimics human lung transplantation, but
because the investigator can collect samples that mimic clinical
investigations of human lung transplant recipients, such as
arterial blood gases (ABGs), bronchioalveolar lavages (BALs),
and lung parenchymal tissue. The most important tests, such as
spirometry and plethysmography have been reported by some
groups (21,50), but are not currently routinely used by us. To
date our laboratory has relied mostly on arterial blood gas (ABG)
measurement of a mouse supported solely by the lung graft after
hilar clamping of the native lung to evaluate function as well
as bronchoalveolar lavage, wet dry ratio, histology, FACS, and
immunohistochemistry. We have focused on these tests as they
mirror the techniques of periopertaive graft assessment used in
the clinics. Some of these techniques are described below.
Arterial Blood Gas measurement(ABG)
(I) Anesthetize the recipient mouse with an intraperitoneal
(i.p) injection of Ketamine (5-6 μg/g) and Xylazine (7-8 μg/g).
This is about 2/3 of the regular dose used during the actual
transplantation procedure as often after transplantation the
recipient cannot tolerate a regular dose of anesthetics (5).
(II) Initiate mechanical ventilation of both lungs with 100%
FiO2 for 4 minutes prior to hilar clamping.
(III) Occlude the hilum of the native non-transplanted lung
(use either a clip or a 6-0 silk tie. Figures 6, 7) and ventilate 4 to
10 more minutes prior to drawing blood.
Figure 6. Occlude the hilum of native non-transplanted lung with clip.
Figure 7. Occlude the hilum of native non-transplanted lung with
6-0 silk.
Here, the microsurgeon needs to detach the right lower
lobe from the esophagus very carefully and free the entire right
lung (in case of left lung transplantation) from connective
tissue without bleeding. The total circulating blood volume in
the mouse is only around 2 mL (6-8% of whole body weight),
and literally any blood loss will affect hemodynamics, result
in circulatory instability and impact the ABG measurement.
Ventilate both lungs for 4 minutes before occluding the native
right lung in order to assess the function of the graft. The
duration of single lung ventilation prior to drawing ABG is controversial and must be tailored based on experimental
conditions (1,26). After transplantation either ischemiareperfusion
injury or graft rejection can cause severe lung graft
dysfunction. Poor oxygenation as a result of this type of injury
can lead to myocardial ischemia, heart failure, and death in
approximately 10 minutes. As mice consume large quantities
of O2 even at rest (31,33), the fall in PaO2 and the associated
oxyghemoglobin desaturation occurs fairly rapidly (51). Thus,
in our experience drawing the ABG after 4 to 10 minutes of
single lung ventilation will allow for a sensitive assessment of
graft function with reliable and reproducible data obtained
by matching the period of single lung ventilation between
experimental and control groups.
(IV) Use a 1mL heparin coated syringe with 25 G needle to
draw blood from the left ventricle or ascending aorta to measure
ABG.
Broncho-alveolar lavage (BAL)
BAL is used to analyze the influx of inflammatory cells into the
airways after human lung transplantation (52) and has been
used by our laboratory to assess lung grafts (3,53). Using current
techniques BAL fluid collection originates from both right and
left lung as the lavage is performed with an open chest while
intubating the main trachea (Figure 8). The advantage of this
technique is that it is easy to perform but the disadvantage is that
it mixes the airway cell infiltrates of the native and transplanted
lung. Nevertheless, our data describe that airway inflammatory
cells, such as neutrophils, correlate with tissue infiltration in
the transplanted lung (4) and thus we have routinely utilized
tracheal BAL to sample the airways and are currently developing
methods to use a longer catheter that can be inserted into either the left or right lung selectively. Alternatively one can occlude
the native lung in order to collect the BAL sample exclusively
from the graft.
Figure 8. The current method for BAL.
Two-photon microscopy
As the lung is constantly exposed to both innocuous and potentially
noxious antigens, a thorough understanding of both innate and
adaptive immune responses in this organ is essential (30). Twophoton
microscopy has evolved into a powerful tool that
can allow for observation of cellular interactions in real time.
Such a technique has allowed us to substantially extend our
understanding of immune responses (54-59). Recently, our
group has expanded 2-photon microscopy to the study of
the inflammatory responses in the lung, which has allowed
us to study ischemia reperfusion injury in vivo (28,55,56).
By relying on this approach we were able to demonstrate
that, contrary to popular belief, monocytes coordinate the
transendothelial migration of neutrophils into inflamed tissue.
We determined that depletion of blood monocytes impairs
neutrophil recruitment to the lung, which could have important
implications for the design of therapeutic strategies to treat
inflammatory lung diseases (28). Based on this experience, we
have expanded the use of intravital two-photon microscopy to
investigate cellular trafficking behavior after lung transplantation.
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Scientific uses of mouse orthotopic lung transplantation
Lung transplantation is an established therapy for a variety of
end-stage pulmonary disease. Importantly, long-term outcome
after lung transplantation are far worse than those of other solid
organs (60). Immunologic and non-immunologic mechanisms
that contribute to acute and chronic graft lung dysfunction
remain poorly understood and the mouse lung transplantation
model presents a unique tool that can allow us to study innate
and adaptive immune responses after lung transplantation.
Ischemia-reperfusion (I-R) injury-mediated primary graft
dysfunction (PGD)
PGD is a form of acute lung injury that results from
inflammatory changes induced by I-R injury (61). PGD is graded
based on PaO2/FiO2 (P/F) ratio and radiographic infiltrates
assessed at several time points up to 72 hours after transplantation
(Table 2). By definition this form of injury arises within the first
72 hours following lung transplantation and is a leading cause
of early morbidity and mortality after transplantation. PGD is
characterized by impaired oxygenation and pulmonary edema
and affects up to 80% of all lung transplant recipients (62-68).
In addition, PGD has been linked to the development of chronic
allograft rejection manifested by bronchiolitis obliterans (69).
Thus, a better knowledge of the pathophysiology of I-R injury
should facilitate a better understanding of PGD. Biomarker
phenotyping should become possible in order to develop
novel therapeutics and reduce the impact of PGD on lung
transplant outcomes (62). Neutrophils and factors that control
their production and activation play a critical role in I-R injury.
Over the last 5 years using the mouse model of orthotopic lung transplantation our group has focused heavily on this cell type.
We have delineated that neutrophils isolated from the airways
of lung transplantation recipients stimulate donor dendritic
cells (DCs) in a contact-dependent fashion to augment their
production of IL-12 and expand alloantigen-specific IFN-γ(+) T
cells. DC IL-12 expression is largely regulated by degranulation
and induced by TNF-α associated with the neutrophil plasma
membrane. Extended cold ischemic graft storage enhances
G-CSF-mediated granulopoiesis and neutrophilic graft
infiltration, resulting in exacerbation of I-R injury after lung
transplantation. I-R injury prevents immunosuppressionmediated
acceptance of mouse lung allografts unless G-CSFmediated
granulopoiesis is inhibited (19). In addition, we also
identified that transcriptional coregulator B cell leukemia/
lymphoma 3 (Bcl3) limits granulopoiesis under inflammatory
conditions. Bcl3-deficient myeloid progenitors demonstrated
an enhanced capacity to proliferate and differentiate into
granulocytes following G-CSF stimulation, whereas the
accumulation of Bcl3 protein attenuated granulopoiesis in an
NF-κB p50-dependent manner (70). Future experiments will
focus on therapeutic strategies to modulate the activation of and
degranulation of neutrophils in order to ameliorate pulmonary
graft injury.
IAcute cellular rejection (ACR)
According to the ISHLT Registry, 36% of lung transplant
recipients experience at least one episode of ACR within
the first year after transplantation (71). Risk factors for ACR
remain poorly defined. The degree of major histocompatibility
antigen (MHC) discordance between donor and recipient has
been identified as a risk factor in some studies (67-69,72). The
mouse model of lung transplantation offers an ideal platform
to study ACR as inbred mouse strains have well defined MHC
antigens allowing for evaluation of ACR in fully mismatched,
minor antigen mismatched, and partially matched (by using F1
crosses as graft donors for example) situations. Since the mouse
lung transplantation model was established, we have verified
histopathologically and flow cytometrically that this new animal
model can recapitulate acute lung allograft rejection successfully
(1). We have also demonstrated that, similar to other organs
(73), treatment of lung allografts with anti-CD28-B7 and CD40-
CD40 Ligand co-stimulatory blockade can prolong allograft
survival indefinitely in several strain combinations (7). Dodd-o
and colleagues found that anti-CD154 antibody therapy alone
is sufficient to attenuate ACR in an MHC mismatched mouse
orthotropic lung transplant model. Improved lung allograft
acceptance in anti-CD154 Ab treated recipients was associated
with abrogated CD8+ and CD4+ allospecific effector responses
and increased frequencies of CD4+CD25+Foxp3+ regulatory
T-cells in the lung allografts (74). Our group has also provided
evidence that pulmonary nonhematopoietic cells, through
their expression of MHC-II, play a critical role in downregulating
CD4+ T cell-mediated immune responses in vivo (2,4). Obviously,
this new mouse model will allow for the design of novel studies
that elucidate mechanisms of ACR and provide rationale for the
development of therapeutic approaches.
Chronic rejection – Obliterative bronchiolitis (OB)
OB is a form of chronic rejection specific to the lung and is
characterized by progressive fibrosis and obliteration of the small
and medium-sized airways of the donor lung (75). The mechanisms
leading to the development of this condition still remain unclear
(76). Although lymphocytes are observed in the bronchial wall, the
prominent cell type found in the BAL is neutrophils (77). Recently,
evidence suggests the involvement of multiple factors such excessive
activation of innate immune responses, abnormal angiogenesis and
failure of appropriate epithelial regeneration and fibroproliferative
tissue remodeling (78). Laboratory experimentation using animal
models forms an important component of a “bench-to-bedsideto-
bench” approach that can both increase our understanding
and lead to the development of novel therapeutic strategies for
this (76).
Of the several different animal OB models available, each
has advantages and limitations. There is not an “ideal” model that precisely reproduces what happens to humans after lung
transplantation. The lesions of OB are thought to represent the
shared histological outcome of injury to the airway epithelium
and subcellular matrix by an array of immune and inflammatory
insults. From a clinical standpoint, OB remains heterogeneous,
varying both in timing of onset after transplantation and
aggressiveness in clinical course (79). So, it is not practical or
scientifically desirable to test multiple contributing factors at
the same time. Pulmonary chronic rejection shares features
of chronic rejection observed in other solid organ. Chronic
rejection in all organs mainly manifests as fibrosis in the graft
resulting in loss of function and eventually grafts loss (80).
There are several immunologic antigens involved in this fibrosis,
including major and minor histocompatibility antigens (81)
as well as self-antigens (82) as both can emerge as targets of
immune responses after transplantation (83-85).
In theory the orthotopic mouse lung transplantation model
has great advantages for studying the pathogenesis of OB as
it offers a physiological model of the human lung transplant
environment. Lung function and BAL can be assessed at
various times, various drugs can be tested for efficacy, and
transgenic and knock-out strains are available to model human
diseases. However, several physiologic factors in the mouse
lung anatomy and physiology create potential obstacles in the
study of OB. The initial OB process starts with a lymphocytic
infiltrate of the sub-mucosa of the airways followed by
the migration of the lymphocytes through the basement
membrane into the epithelium (86). At this site, epithelial cell
necrosis occurs with denudation of mucosa. In the mouse lung,
there is lack of submucosal glands in the bronchioles, but a
high numbers of locally resident Clara cells. One of the main
functions of Clara cells is to protect the bronchiolar epithelium
from injury. They accomplish this by secreting a variety of
proteins, including Clara cell secretory protein (CCSP) and a
solution similar to lung surfactant. They are also responsible
for detoxifying inhaled harmful substances. Clara cells also act
as a stem cell and multiply to differentiate into ciliated cells
that can regenerate bronchiolar epithelium. This is a possible
reason why mouse airway epithelium remains intact and fully
differentiated in lung allografts, despite profound vascular
rejection (7). Since 2007, a few separate groups have tried to
establish an OB model in the mouse. Although two groups
have found OB lesions by histology, the experimental design
and the histological features still need further clarification
(9,27). Currently we are focusing on three factors that may
influence the development of OB including: (I) donor and
recipient strain combinations; (II) time period necessary
to see chronic rejection in the mouse and; (III) the form of
immunosuppression that may allow for OB to develop. Once
established and reproducible, such a model could hold great
promise for further mechanistic studies and may be used to
accelerate the development of new strategies for the prevention
or treatment of pulmonary chronic rejection.
Studies of Non-hematopoietic stromal cells
While bone marrow-derived hematopoietic cells, such as
dendritic cells, play a critical role in pulmonary physiology
(87-92), we as well as others have focused on the contribution
of non-hematopoietic cells in pulmonary immune responses
(4,93). Traditional methods for separating the physiology
of hematopoietic from non-hematopoietic cells involves the
creation of bone marrow chimeras by lethal irradiation of a
recipient mouse followed by reconstitution with bone marrow
from a mutant strain (94). Donor irradiation, however, may
change the physiology of non-hematopoietic cells and bone
marrow chimeras can also suffer from autoimmune disease (4).
We have recently demonstrated that the transplantation of a
left lung into a congenic host leads to the rapid substitution
of donor-derived hematopoietic cells with those of the host,
leading to the creation of a “chimeric lung graft” in an otherwise
immunocompetent host. Furthermore, such substitution occurs
for multiple types of hematopoietic cells such as T cells (4,5)
and dendritic cells (Figure 9). Such a model can thus be used
to study both immunology and physiology of pulmonary nonhematopoietic
cells in the absence of irradiation.
Figure 9. Rapid substitution of professional antigen presenting cells,
such as dendritic cells, occurs after orthotopic lung transplantation.
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Conclusions
Although technically difficult, orthotopic lung transplantation
in the mouse is feasible and reproducible. It has been used for
multiple scientific avenues of investigation for the last 5 years by
our laboratory as well as others. Obviously, it is important for
researchers to understand the physiology behind this model as well
as its advantages and limitations. Well-designed mouse orthotopic
lung transplantation experiments provide a great opportunity to
study mechanisms that affect lung allograft survival and explore
new therapies for a wide variety of human lung diseases. | ||||||||||||||||||||||||||||||||||||
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Acknowledgements
Supported by American Thoracic Society/Lungevity Foundation,
NIH 1R01HL094601, the Barnes Jewish Research Foundation,
The American Association for Thoracic Surgery and the Thoracic
Surgery Foundation for Research and Education.
Disclosure: The authors declare no conflict of interest.
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References
Cite this article as: Lin X, Li W, Lai J, Okazaki M,
Sugimoto S, Yamamoto S, Wang X, Gelman AE,
Kreisel D, Krupnick AS. Five-year update on the
mouse model of orthotopic lung transplantation:
Scientific uses, tricks of the trade, and tips for success.
J Thorac Dis 2012;4(3):247-258. doi: 10.3978/
j.issn.2072-1439.2012.06.02
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