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Mouse Embryonic Diastema Region Is an Ideal Site for theDevelopment of Ectopically Transplanted Tooth GermYiqiang Song,

1Mingquan Yan,

1Ken Muneoka,

1and YiPing Chen

1,2*

1Department of Cell and Molecular Biology, Tulane University, New Orleans, Louisiana

2Department of Oral Biology, Ohio State University Health Sciences Center, Columbus, Ohio

*Correspondence to: YiPing Chen, Department of Oral Biology, Ohio State University College of Dentistry, 305W. 12th Avenue, Columbus, OH 43210. Email: [email protected] Dr. Songs present address is Department of Oral Biology, University of Illinois at Chicago College of

Dentistry, 801 S. Paulina Street, Chicago, IL 60612.The publisher's final edited version of this article is available free at Dev DynSee other articles in PMC that cite the published article.

y Other Sections o AbstractINTRODUCTIONRESULTS AND DISCUSSIONEXPERIMENTAL

PROCEDURESReferencesAbstractThe anterior eye chamber and the kidney capsule of the mouse have been traditionally used

for long-term culture of tooth germ grafts. However, although these sites provide an

excellent growth environment, they do not represent real in situ sites for the development of

a grafted tooth germ. Here, we describe a protocol to transplant a tooth germ into the

mandibular diastema region of mouse embryos using exo utero surgery. Our results

demonstrate that the mouse embryonic diastema region represents a normal physiological

environment for the development of transplanted tooth germs. Transplanted tooth germs

developed synchronically with and became indistinguishable from the endogenous ones.

These ectopic teeth were vascularized and surrounded with nerve fibers, and were able to

erupt normally. Thus, the exo utero transplantation approach will provide a new avenue to

study tooth development and regeneration.

Keywords: tooth germ, tooth development, exo utero surgery, diastema


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PROCEDURESReferencesINTRODUCTIONGiven the relatively concise morphology and the ease with which it can be accessed,

manipulated, cultured, and grafted ex vivo, the mouse tooth germ has long been an

excellent model to study vertebrate organogenesis (Thesleff et al., 1996). Several

approaches have been developed to study tooth development ex vivo. The in vitro organ

culture has been routinely used, in which tooth germs are cultured in the Trowell type organ

culture dish (Thesleff et al., 1996). This method provides an easy and quick culture for

short-term growth of tooth germ and also provides an openly accessible environment for

the manipulation of tooth germ such as addition of growth factors. However, the artificial

culture condition limits not only the length of culture time but also the shape and size of

explanted tooth germs. The other method that has been used extensively for long-term tooth

germ culture is the kidney capsule culture in which tooth germs or organ primordial tissue

are grafted and cultured underneath the kidney capsule of adult mice (Morio, 1985).

Grafted tooth germs can develop into a well-differentiated tooth (Zhang et al., 2003b).

Actually, many parts of mouse body have been tested for hosting grafted tooth germs.

These sites include the anterior eye chamber (Kollar and Fisher, 1980; Yoshikawa and

Kollar, 1981; Lumsden and Buchanan, 1986), renal subcapsular site (Bartlett and Reade,

1973), subcutaneous tissue (Ivanyi, 1965), cheek pouch (Al-T

alabani and Smith, 1979), andspleen (Ishizeki et al., 1987). In addition, chick embryonic wing buds were also shown to

be an effective in vivo culture system for tooth development (Koyama et al., 2002).

However, even though some of these models provide an excellent growth environment for

tooth grafts, they represent a developmental and physiological environment that is distinct

from the oral cavity where tooth development normally occurs.

The best and ideal site for ectopic tooth growth is within the oral cavity. The oral cavity of

the adult mouse has been successfully used to support the development of grafted tooth

germ (Ohazama et al., 2004; Nakao et al., 2007). However, the mouse after birth is full of

activity like daily food intake, which may influence and exert mechanical pressure on the

normal development of grafted tooth germ. It would be a better choice to nurture a tooth

germ in the oral cavity of the mouse at a comparable developmental stage. In mice, a

toothless region called diastema exists between the incisor and the first molar. This region


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is believed to be filled with incisor, canine, and premolars before their disappearance

during the rodent evolution (Luckett, 1985; Peterkova et al., 1995). Today, the rudimentary

tooth germs still can be observed in the upper diastema of the mouse or sibling vole

embryos, and then these tooth germs are removed apoptotically through an unknown

mechanism (Keranen et al., 1999). On the other hand, the initiation of tooth development in

the mandibular diastema region is evidenced by a weak expression ofShh just anterior to

the first molar primordia (Mustonen et al., 2004). Accordingly, disturbance of the normal

gene regulation network in this area may results in an extra tooth (Mustonen et al., 2003;

Zhang et al., 2003a; Tucker et al., 2004; Kassai et al., 2005). Based on these observations,

we reasoned that the mandibular diastema region could be an ideal site for the development

of a tooth germ in addition to the endogenous tooth forming loci. To test this hypothesis,

we carried out an exo utero surgery to transplant tooth germ. Here, we describe this method

and demonstrate that transplanted tooth germs develop normally in the mouse embryonic

diastema region, indistinguishable from the endogenous ones.


PROCEDURESReferencesRESULTS AND DISCUSSIONTransplantation of Tooth Germ to the Mandibular Diastema of Mouse Embryo

by Exo Utero Surgery

One of the greatest advantages of using avian embryo as a developmental biology model is

the accessibility of its embryos throughout the entire developmental process. In the mouse,

however, accessing to the developing embryo is always difficult and poses a great

challenge due to the complex architecture of the mouse gestation organ. The finding that

mouse embryos can develop normally outside the uterine myometrium has substantially

extended the accessibility of mouse embryo to a much earlier stage, and moreover, such an

exo utero mouse embryo can survive some surgical manipulations to term (Muneoka et al.,

1986a,b). Benefiting from these findings, exo utero surgery has evolved to become a

powerful tool to study mouse embryonic development (Ngo-Muller and Muneoka, 2000). It

has been widely used in studies of nerve system development such as migration of the

neural crest cells during early mouse embryogenesis (Serbedzija et al., 1992), limb

development and regeneration (Han et al., 2003), as well as in teratology research where


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effects of chemicals on the developmental mouse embryos can be studied. A recent review

has summarized the applications of mouse exo utero development system in biological

studies (Hatta et al., 2004).

We were originally seeking to identify an ideal site for the best development and growth of

ectopically grafted tooth germ or recombinant tooth germ after genetic modification in vitro

(Song et al., 2006). Compared with the kidney capsule, the anterior eye chamber, or other

ectopic loci, the oral cavity of the mouse appears to represent the most natural environment

for tooth development. The diastema, a toothless region between the incisor and molar, has

been demonstrated to host a full array of teeth before they were evolutionally lost. Teeth

indeed develop in the mandibular diastema of some genetically modified mice (Mustonen

et al., 2003; Tucker et al., 2004; Zhang et al., 2003a; Kassai et al., 2005). We, therefore,

deduced that the mandibular diastema of the mouse embryo had an intrinsic ability to

support tooth development.

Although exo utero surgery can be carried out on mouse embryos as early as embryonic

day (E) 11.5, a better survival rate is achieved with surgery on E13.5 or older embryos

(Muneoka et al., 1986a). On the other hand, tooth germs of E13.5 mouse embryos are at the

bud stage, which provides an ideal size and solidity for surgical operation compared with

that from earlier stages. Thus, we began with transplanting E13.5 mouse tooth germs into

the mandibular diastema of E13.5 host embryos. Such an arrangement allows the

endogenous tooth primordia to be the control, which provides standard developmental

indexes side by side for the evaluation of the ectopic tooth growth. Here, we further took

advantage of tooth germs from Rosa26-Egfp mouse embryos, of which tissue can be easily

detected and distinguished from the host embryo under the fluorescent microscope by

activation through fluorescein isothiocyanate (FITC) -filter set.

For exo utero surgery, the incisor tooth germ was isolated from the E13.5 Rosa26-Gfp

mouse embryo. At this stage, the incisor primordia is much better for transplantation

experiments due to their smaller size compared with the molar tooth germs. It can be easily

grafted into a small incision made in the diastema region of host embryos. As described in

detail in the Experimental Procedures section, the same stage embryos were properly

exposed from the uterus of an anesthetized female mouse. As shown in Figure 1A, an E13.5

host embryo was exposed after opening the uterus and removing the extraembryonic


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membrane in a timed-pregnant mouse. Under adequate illumination, details of the embryo

can be easily defined under the dissection microscope. Before grafting into the small

incision made in the diastema region, the isolated incisor germ was carried on a thin copper

wire to the embryonic mandible. The tooth germ was then carefully placed into the incision

with the epithelial side face up. By piercing the neighboring host tissue with the copper

wire, the transplanted tooth germ was fixed at the incision site. Figure 1B shows a

transplanted Rosa26-Egfp incisor primordia fixed at the mandibular diastema region of an

E13.5 wild-type host embryo. Under the activation light through the FITC-filter set, the

ectopic tooth germ was emanating green fluorescent light.

Fig. 1 Ex utero surgery for tooth germ transplantation into the mandibular diastema of

embryonic day (E) 13.5 mouse embryo. A: An E13.5 mouse embryo is exposed after

opening of the uterus and extraembryonic membrane. B: An E13.5 host embryoreceives an incisor (more ...)

Fig. 1Ex utero surgery for tooth germ transplantation into the mandibular diastema of embryonic day (E) 13.5

mouse embryo. A: An E13.5 mouse embryo is exposed after opening of the uterus and extraembryonicmembrane. B: An E13.5 host embryo receives an incisor tooth germ from E13.5 Rosa26-Egfp embryo in

the mandibular diastema region. The grafted tooth germ that was fixed by two copper wire pins showed


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fluorescence under the fluorescent microscope. C: A full-term host mouse carrying the grafted tooth

germ, as indicated by fluorescence. AM, amnion membrane; CB, cotton ball; CW, copper wire pin; E,

eye; FL, front limb; GT, GFP mouse embryo derived tooth germ; H, head; MD, mandible; MX,maxillary.

Transplanted Tooth Germ Develops Normally in the Diastema

Histological analysis of a full-term host embryo shows a well-developed tooth germ in the

transplanted site (Fig. 2A). Both epithelial and mesenchymal structures of the transplanted

incisor are comparable to that of the endogenous incisors (Fig. 2B). To confirm the

supportive role of host tissue in the ectopic tooth development, we further examined blood

vessel invasion and innervation of the transplanted tooth. By immunohistochemical staining

using antibody against laminin, a molecular marker for blood vessel endothelial cells, we

were able to detect blood vessels in the dental pulp of both the endogenous and the

transplanted teeth (Fig. 2C,D), clearly demonstrating the presence of blood vessels in thetransplanted tooth. The invasion of blood vessels provides the transplanted tooth with

nutrients through the blood supplies from the host. Moreover, using antibodies against

neurofilament, we have detected nervous termini around the transplanted tooth, indicating

that the transplanted tooth will have the similar potential as the endogenous teeth to be

innervated (Fig. 2E,F). Thus, the transplanted tooth germ develops synchronically with the

endogenous tooth in the host embryo. Based on the facts that blood vessels are not present

in the developing mouse tooth before the cap stage (E14.5; Luukko et al., 2005) and that

tooth innervation does not occur until birth (Tsuzuki and Kitamura, 1991; Kettunen et al.,

2005), we conclude that these blood vessel cells and neuron cells within or surrounding the

transplanted tooth are derived from the host.

Fig. 2 Development of transplanted tooth germ in the term host mouse. A: A

coronal section through a host mouse head shows a grafted incisorgerm (GI) in the mandibular diastema adjacent to the endogenous

incisor germ (EI). B: Higher magnification of A shows (more ...)


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Fig. 2Development of transplanted tooth germ in the term host mouse. A: A coronal section through a hostmouse head shows a grafted incisor germ (GI) in the mandibular diastema adjacent to the endogenous

incisor germ (EI). B: Higher magnification of A shows a well developed ectopic incisor (arrow),

comparable to the adjacent endogenous one. C,D: Innmunohistochemical staining shows cells positive

for laminin, a molecular marker for blood vessel endothelial cells, in the dental pulp of a transplantedincisor in a host mouse at birth (C), compared with the endogenous incisor at the same age (D). E,F:

Immunostaining for neurofilament shows nerve termini (arrow) around a transplanted incisor (E), and in

the endogenous control tooth (F). AB, alveolar bone; AM, ameloblast; BV, blood vessel; D, dentin; DP,dental pulp; EI, endogenous incisor; GI, grafted incisor; OB, odontoblast; T, tongue; N, nerve.

Of interest, when E13.5 embryos were used as hosts for tooth transplantation, all the hosts

(21 in total) that were monitored died within 1 day after caesarian section. Examination of

gross morphology identified the cleft palate phenotype in all the surgical embryos,

explaining the postnatal lethality. It is known that the palate shelves elevated from a

perpendicular to the horizon position and fused to each other above the tongue between

E13.5 and E14.5. Therefore, our surgical operation in the oral cavity of E13.5 embryo must

have disturbed such an elevation and/or fusion process of the palate shelves, resulting in the

cleft palate formation.

Because the palate shelves have already fused in E14.5 mouse embryos, we decided to use

E14.5 mouse embryos as host for tooth transplantation, in hope of getting living pups with

an ectopic tooth in their mouth. As expected, none of the pups (25 in total) that received

tooth germ transplantation at E14.5 developed a cleft palate phenotype. The mice were fed

by a foster mother immediately after C-section, and grew up as normal mice. Figure 3


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shows such a weaned mouse carrying an ectopic incisor growth in the mandibular diastema.

These results clearly demonstrate that the transplanted tooth germ can develop normally in

the diastema region of mouse embryo and can eventually erupt and form as a normal tooth

in the host mouse oral cavity.

Fig. 3 Growth of a transplanted incisor in the mandibular diastema of a hostmouse. The grafted tooth erupted and grew at the comparable size as the

endogenous ones. LI, lower incisor; GI, grafted incisor; UI, upperincisor; N, nasal

Fig. 3Growth of a transplanted incisor in the mandibular diastema of a host mouse. The grafted tooth erupted

and grew at the comparable size as the endogenous ones. LI, lower incisor; GI, grafted incisor; UI, upperincisor; N, nasal

The mechanism of how some vertebrates lost their teeth during evolution remains elusive.

It could be a result of loss of gene function that is critical for tooth development. In avian,

the remnant of developing tooth primordia can be found in the oral cavity too, and the

abortion of tooth development is at least partially due to the loss of odontogenic Bmp4

expression (Chen et al., 2000). On the other hand, tooth agenesis could also be a result of

an active inhibitory mechanism that is present in the oral cavity to repress tooth

development. It is evidenced that Gas1, a diffusible protein acting as Shh antagonist, is able

to sequester Shh and, therefore, to block Shh function in the mouse diastema region. Such

an antagonistic mechanism may be used to delineate the border of tooth forming sites and

the diastema region at the beginning of tooth development, or required to constantly inhibit

tooth development within the diastema (Cobourne et al., 2004). Nevertheless, we show here

that, at or after E13.5, the mandibular diastema of mouse embryo serves as an excellent site


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to support tooth development. Our results suggest that the grafted tooth germ is able to

override any inhibitory mechanism or can develop independent of the inhibitory

mechanism. Alternatively, the constant inhibitory mechanism discussed above may have

disappeared in the diastema region at the time when tooth germ is grafted.

Similar to many other ectopic sites in living animals, particularly the mouse kidney capsule,

the embryonic diastema region appears to be excellent for grafted tooth development and

differentiation. However, the approach we describe here also has several advantages, such

as (1) it makes it possible to study tooth eruption of embryonic lethal mutant mice, and (2)

it permits studies on tissue interactions between grafted tooth germ and the host supportive

tissues in the oral cavity. On the other hand, this approach has its limitation as well. The

size of a graft is a concern. For example, an E13.5 molar germ was proven too big to be

grafted properly. In conclusion, while the diastema region of the mouse mandible does not

grow endogenous teeth, it does fully support tooth formation when ectopic tooth germ is

grafted.


PROCEDURESReferencesEXPERIMENTAL PROCEDURESTooth Germ Preparation and Transplantation by Means of Exo Utero Surgery

Tooth germs at the bud stage were used as donors in this study. Embryos at E13.5 were

collected from timed pregnant CD-1 orRosa26-Egfp mice. The incisor tooth germs were

isolated from the mandibles and kept in ice-cold PBS for use. A fine copper pin was used to

pierce through each tooth germ at the mesenchyme, making it easier to transfer the tooth

germ from the dish to the surgical embryo with forceps. Exo utero surgery was performed

as described previously (Ngo-Muller and Muneoka, 2000). In this study, we used E13.5 and

E14.5 embryos as recipients for tissue transplantation. Briefly, timed pregnant CD-1 mice

at 13.5 or 14.5 gestation were anesthetized by intraperitoneal injection of Nembutal at a

dose of 10 l/g body weight. After properly anesthetized, the mouse was placed on a

surgical platform. The skin and body wall of the abdomen were sequentially opened by

scissors and forceps. The uterus was subsequently opened with iridectomy scissors at the

opposite of placental side in a continuous longitudinal incision. By rolling a dry cotton-

tipped applicator between the placenta and the uterus, the unwanted embryos were removed


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with minimal bleeding from the uterus. Usually, four embryos were left in the abdomen of

each female for grafting. Blood and debris were washed away with lactated Ringers

solution before further operation. Extra care needs to be taken to avoid damaging the major

blood vessels in the extraembryonic membrane when an incision was made in the

membrane. The head of the host embryo was allowed to pop out through the incision on the

extraembryonic membrane, and then was properly positioned with the help of sterile cotton

balls as cushions. A small slit was made by fine forceps within the diastema region of the

hosts mandible. Donor tooth germ was inserted into the incision with the dental epithelial

side facing the oral cavity, and then was fixed by fine copper pins. After completion of

tissue transplantation, the extraembryonic membrane was closed by suturing. The abdomen

cavity was then sutured after cotton balls were removed and the abdomen cavity was totally

cleaned with lactated Ringers solution. Typically, the mouse wakes up within 1 hr after

surgery. A warm heating pad was used to help the mouse maintain body temperature during

this process.

The pregnant female mouse was housed and closely monitored in an isolated cage until

term. To collect experimental embryos, the host female mouse was killed, and surgical pups

were collected through C-section. For further development purposes, the operated embryos

were stimulated to begin breathing by rubbing cotton-tipped applicator on their tail tips.

Any fluids in their mouth were also cleaned with a cotton-tipped applicator. When they

were breathing regularly and moving about, the experimental newborns were transferred to

the cage with a nursing mother that had delivered her litter at the same day or 1 day before.

An animal protocol for the exo utero surgery was approved by the IACUC ofTulane

University. The entire surgery procedure strictly followed the animal protocol.

Histology and Azan Dichromic Staining

The recipient mice were killed at birth. The heads were harvested and fixed in 4%

paraformaldehyde (PFA)/ PBS, de-calcified by demineralization in 0.1 M

ethylenediaminetetraacetic acid/PBS for a week with several changes of solution. Samples

were then dehydrated through graded ethanol, cleared with xylene, embedded in paraffin,

and sectioned at 10 m. Sections were processed with Azan di-chromic staining, as

described previously (Presnell and Schreibman, 1997).


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Immunohistochemistry

The recipient mice were collected at the delivery day, and the heads were removed and

processed for frozen sections. OCT-embedded samples were sectioned at 10 m, and the

cryostat sections were fixed for 30 min at room temperature with 4% PFA and analyzed by

indirect immunohistochemical staining. The rabbit anti-mouse laminin antibody

(Chemicon, Temecula, CA) was used as the primary antibody specific for blood vessel

endothelial cells, and the rabbit anti-mouse neurofilament 200 antibody (Sigma, St. Louis,

MO) was used as the primary antibody for nerve cells. The horseradish peroxidase-coupled

goat anti-rabbit IgG were used as the secondary antibody (Sigma). Immunohistochemical

staining was performed according to the protocol provided with the antibodies by the

manufacturers.

AcknowledgmentsY.P.C. and K.M. were supported by the NIH.

Grant sponsor: the NIH; Grant number: DE15123; Grant number: DE16623; Grant number:

HD43277.


PROCEDURESReferencesReferences

1. Al-Talabani NG, Smith CJ. A histological study of the effect of developmental stage of tooth-germ on

transplantation to hamster cheek pouch. Arch OralBiol.1979;24:933937. [PubMed]

2. Bartlett PF, Reade PC. The antigenicity of mouse tooth germs. I. Isogenic and allogenic transplantation.

Transplantation.1973;16:479488. [PubMed]

3. Chen YP, Zhang Y, Jiang TX, Barlow AJ, St Amand TR, Hu Y, Heaney S, Francis-West P, Chuong CM,

Maas R. Conservation of early odontogenic signaling pathways in Aves. Proc NatlAcadSci U SA.

2000;97:1004410049. [PMC free article] [PubMed]

4. Cobourne MT, Miletich I, Sharpe PT. Restriction of sonic hedgehog signalling during early tooth

development. Development.2004;131:28752885. [PubMed]

5. Han MJ, Yang XD, Farrington JE, Muneoka K. Digit regeneration is regulated by Msx1 and Bmp4 in fetal

mice. Development.2003;130:51235132. [PubMed]


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Stem/Progenitor CellMediated De Novo Regeneration ofDental

Pulp with Newly Deposited Continuous Layer ofDentin in an In

Vivo ModelGeorge T.-J. Huang, D.D.S., M.S.D., D.Sc.,

1,2,3Takayoshi Yamaza, D.D.S., Ph.D.,4,*

Lonnie D. Shea, Ph.D.,5

Farida Djouad, Ph.D.,3

Nastaran Z. Kuhn, Ph.D.,3

Rocky S. Tuan,

Ph.D.,3, and Songtao Shi, D.D.S., Ph.D.41Section of Oral and Diagnostic Sciences, Division of Endodontics, College of Dental Medicine, Columbia University,New York, New York.2Department of Endodontics, Prosthodontics, and Operative Dentistry, College of Dental Surgery, Dental School,University of Maryland, Baltimore, Maryland.3Cartilage Biology and Orthopaedics Branch, National Institute of Arthritis, and Musculoskeletal and Skin Diseases,National Institutes of Health, Department of Health and Human Services, Bethesda, Maryland.4Center for Craniofacial Molecular Biology, University of Southern California School of Dentistry, Los Angeles,California.5Department of Chemical and Biological Engineering, McCormick School of Engineering, Northwestern University,Evanston, Illinois.

Corresponding author.Address correspondence to: GeorgeT.-J. Huang,D.D.S.,M.S.D.,D.Sc.,DepartmentofEndodontics,Boston University,HenryM.Goldman SchoolofDentalMedicine,100E. Newton St.,Boston,MA02118.E-mail:Email: [email protected] *

Present address: Department of Oral Anatomy and Cell Biology, Kyushu University Graduate School of Dental Science,Fukuoka, Japan.Present address: Department of Orthopaedic Surgery and Center for Celluar and Molecular Engineering, University ofPittsburgh School of Medicine, Pittsburgh, Pennsylvania.Received July 25, 2009; Accepted September 8, 2009.

This article has been cited by other articles in PMC.

y Other Sections o AbstractIntroductionMaterials and

MethodsResultsDiscussionReferencesAbstractThe ultimate goal of this study is to regenerate lost dental pulp and dentin via

stem/progenitor cellbased approaches and tissue engineering technologies. In this study,we tested the possibility of regenerating vascularized human dental pulp in emptied root

canal space and producing new dentin on existing dentinal walls using a stem/progenitor

cellmediated approach with a human root fragment and an immunocompromised mouse

model. Stem/progenitor cells from apical papilla and dental pulp stem cells were isolated,

characterized, seeded onto synthetic scaffolds consisting of poly-D,L-lactide/glycolide,

inserted into the tooth fragments, and transplanted into mice. Our results showed that the

root canal space was filled entirely by a pulp-like tissue with well-established vascularity.

In addition, a continuous layer of dentin-like tissue was deposited onto the canal dentinal

wall. This dentin-like structure appeared to be produced by a layer of newly formed

odontoblast-like cells expressing dentin sialophosphoprotein, bone sialoprotein, alkaline

phosphatase, and CD105. The cells in regenerated pulp-like tissue reacted positively to

anti-human mitochondria antibodies, indicating their human origin. This study provides the


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first evidence showing that pulp-like tissue can be regenerated denovo in emptied root

canal space by stem cells from apical papilla and dental pulp stem cells that give rise to

odontoblast-like cells producing dentin-like tissue on existing dentinal walls.


MethodsResultsDiscussionReferencesIntroductionREGENERATION OF dental pulp/dentin tissues in the pulp space of teeth serves the ultimate goal

of preserving teeth via endodontic approaches. Attempts to induce tissue regeneration in the

pulp space have been a longstanding quest. Pulp tissue regeneration has been explored

using the biodegradable synthetic material polyglycolic acid seeded with pulp cells, and

results showed pulp-like tissue formation in both invitro and invivo models.13

These

previous approaches were the proof-of-principle studies that only tested the formation of apulp-like soft tissue without dentin. From a clinical perspective, the following issues must

be considered when attempting to regenerate functional pulp/dentin tissues in a root canal

space: (i) regenerated pulp tissue must be vascularized although the blood supply is only

available from the apical end; (ii) newly differentiated odontoblasts should form on the

existing dentinal wall in the root canal space, and (iii) new dentin should be produced by

the new odontoblasts onto the existing dentin.4,5

Using a tooth slice model (horizontal section, 1 mm thick), it was shown that the stem cells

from human exfoliated deciduous teeth seeded onto the synthetic scaffolds that were fabri-

cated in the pulp chamber space formed well-vascularized pulp-like tissue in an invivo

study model. In addition, odontoblast-like cells derived from the pulp-like tissue were

localized against the existing dentin surface.6Tooth slice model was used to ensure blood

supply to the stem cellseeded scaffolds. To date, there is a lack of evidence demonstrating

that the human pulp tissue can be regenerated in an emptied root canal space with only one

opening to the blood supply, and showing that the regenerated pulp tissue would form a

continuous layer of newly deposited dentin onto the existing dentinal walls. Using human

DPSCs from permanent teeth seeded onto a dentin surface, small amounts of discontinuous

dentin-like mineralized tissue on the existing dentin surface have been observed invivo.7

Previously, we reported that the developing organ at the apex of the tooth, the apical

papilla, contained multipotent stem cells, named stem cells from apical papilla (SCAP), that

were able to form pieces of pulp-dentin complex in an animal model.8,9

Moreover, SCAP


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are cells from a developing tissue and are more robust than DPSCs in terms of population

doubling capacity, proliferation rate, telomerase activity, and cell migration ability.8 We

hypothesized that vascularized pulp-like and dentin-like tissues can be regenerated in an

emptied root canal space with DPSCs and SCAP. Using a model system that includes ex

vivoexpanded human DPSCs and SCAP, synthetic scaffolds, human root segments, and

immunocompromised mice, we show here for the first time the regeneration of vascularized

pulp-like tissue and the formation of dentin-like mineral structures depositing onto the

existing dentinal wall in the root canal space.


MethodsResultsDiscussionReferencesMaterials and MethodsSample collec

tion

Normal human-impacted third molars (n = 12) with or without immature roots were

collected from healthy consenting patients (eight donors aged 1624 years) in the Dental

Clinics at the University of Maryland and the University of Southern California. Freshly

extracted teeth were stored in serum-free cell culture medium and transported to the

laboratory for processing. Bone marrow was obtained from the long bone of healthy

consenting patients (aged 2063) undergoing orthopedic surgery at the Walter Reed Army

Medical Center. Sample collection conformed to approved protocols by the respective

Medical Institutional Review Boards and the National Institutes of Health.

Cell culture

Isolation of DPSCs and SCAP was described previously.811

In brief, pulp tissue and apical

papilla were removed from teeth, minced, and digested in a solution of 3 mg/mL

collagenase type I and 4 mg/mL dispase (Sigma-Aldrich, St. Louis, MO) for 3060 min at

37C. The digested mixtures were passed through a 70-m cell strainer (Falcon; BD

Labware, Franklin Lakes, NJ) to obtain single-cell suspensions. Cells were seeded onto six-

well plates and cultured with -minimum essential medium (GibcoInvitrogen, Carlsbad,

CA) supplemented with 1520% fetal bovine serum (FBS; Gemini Bio-Products,

Woodland, CA), 2 mM L-glutamine, 100 M L-ascorbic acid-2-phospate, 100 U/mL

penicillin-G, 100 g/mL streptomycin, and 0.25 g/mL fungizone (Gemini Bio-Products)

and maintained in 5% CO2 at 37C. Colony formation units of fibroblastic cells were

normally observed within 12 weeks after cell seeding and were passaged at 1:3 ratio when


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they reached ~80% confluence. Heterogeneous populations of DPSCs and SCAP were

frozen and stored in liquid nitrogen at passages 02. Cells were thawed and expanded for

experimentation at passage 3. Human bone marrow mesenchymal stem cells (BMMSCs)

were isolated and cultured as previously described1113 and were cultured in expansion

medium containing Dulbecco's modified Eagle's medium and 10% preselected FBS.

Heterogeneous populations of isolated colony formation units of fibroblastic cells were

used for all the assays and tissue regeneration experiments.

Flow cytometry

Cell aliquots of SCAP and DPSCs (>2 105

cells) were washed and resuspended in

phosphate-buffered saline (PBS) + 0.1% FBS, containing saturating concentrations (1:100

dilution) of the following conjugated mouse IgG1,anti-human monoclonal antibodies (BD

Biosciences, San Jose, CA): CD14-PE, CD34-PE, CD45-FITC, CD73-PE, CD90-FITC,

and CD105-PE for 1 h at 4C. Cell suspensions were washed twice and resuspended in

0.1% FBS/PBS for analysis on a flow cytometer (FACS Calibur; BD Biosciences) using

the CellQuest Pro software (BD Biosciences).

Multiple lineage differentiation

Odonto/osteogenicdifferentiation

Cells were seeded onto 48-well plates, grown to ~70% confluence, and incubated in the

differentiation medium containing 10 nM dexamethasone, 10 mM -glycerophosphate, 50

g/mL ascorbate phosphate, 10 nM 1, 25 dihydroxyvitamin D3, and 10% FBS for 5 weeks.

Cultures were fixed in 60% isopropanol, and mineralization of extracellular matrix stained

with 1% Alizarin Red S.

Adipogenicdifferentiation

Cells were seeded onto 12-well plates, grown to subconfluence, and incubated in the

adipogenic medium containing 1 M dexamethasone, 1 g/mL insulin, 0.5 mM 3-isobutyl-

1-methylxanthine, and 10% FBS for 6 weeks. Cells were fixed in 10% formalin for 60 min,

washed with 70% ethanol, and lipid droplets were stained with 2% (w/v) Oil Red O reagent

for 5 min and washed with water.

Neurogenicdifferentiation

Cells at subconfluence in chamber slides were incubated in the neurogenic induction

medium Neurobasal A (Gibco-Invitrogen) with B27 supplement (GIBCO-BRL), 20 ng/mL


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epidermal growth factor (EGF) (BD Biosciences), and 40 ng/mL fibroblast growth factor

(FGF) (BD Biosciences). After 4 weeks, cells were analyzed by immunocytofluorescence

for the expression of the neural cell marker, III-tubulin. Isotype-matched antibodies were

used as negative controls. After fixation in 100% ice-cold methanol, cells were incubated in

blocking buffer (32.5 mM NaCl, 3.3 mM Na2HPO4, 0.76mM KH2PO4, 1.9 mM NaN3,

0.1% [w/v] bovine serum albumin, 0.2% [v/v] Triton-X 100, 0.05% [v/v] Tween 20, and

5% goat serum) for 30 min followed by the addition of monoclonal mouse anti-human III-

tubulin antibodies (Promega, Madison, WI) for 1 h at room temperature. After washing,

cultures were incubated with anti-mouse Alexa Fluor 594 for 1 h at room temperature and

the cell nuclei stained with 4',6-diamidino-2-phenylindole dihydrochloride (DAPI)

(Invitrogen, Carlsbad, CA). Images were analyzed under a fluorescence microscope.

Myogenicdifferentiation

Cells were seeded onto 12-well plates, grown to subconfluence, and induced in the

myogenic medium containing 0.1 M dexamethasone, 50 M hydrocortisone, and 5% horse

serum for 6 weeks. Cells were then harvested for RNA isolation using RNeasy Mini Kit

(Qiagen, Valenicia, CA). cDNA was synthesized with SuperScript reverse transcription

(RT)-polymerase chain reaction kit (Invitrogen) followed by polymerase chain reaction

using Taq DNA polymerase (Invitrogen) to detect the expression of myogenic genes. The

following genes were examined with specific primers: MyoD1 (forward, 5-AAG CGA

CCT CTC TTG AGGTA-3; reverse, 5-GCG CCTTTA TTTTGA TCA CC-3),

myogenin (forward, 5-TAA GGTGTGTAA GAGGAA GTC G-3; reverse, 5-CCA CAG

ACA CAT CTT CCA CTGT-3), myosin heavypolypeptide1 (forward, 5-TGTGAA TGC

CAA ATGTGC TT-3; reverse, 5- GTGGAG CTGGGT ATC CTTGA-3), and GAPDH

(forward, 5-CAA GGC TGA GAA CGGGAA GC-3; reverse, 5-AGGGGG CAG AGA

TGA TGA CC-3).

Scaffold fabrication

A copolymer of poly-D,L-lactide and glycolide (PLG) (75:25 molar ratio) (Boehringer

Ingelheim, Ingelheim, Germany) was employed to create porous scaffolds using a gas

foaming/particulate leaching process, and cells were seeded onto the scaffolds according to

the previously described procedures.14

Briefly, porous polymer scaffolds were formed by

mixing PLG microspheres and salt crystals (diameter 250425 m), which were then


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loaded into a cylindrical mold of 5 (diameter) 2 (height) mm. The mixture was then

compressed at 1500 psi, yielding solid disks and foamed in a pressure vessel using CO2 at

850 psi. Subsequent solvent leaching rendered the scaffolds porous with pore diameters of

250425m.

In vitroanalysis of dental stem cells grown in scaffolds

Each PLG scaffold disk was cut into small pieces of ~1.52.0 mm3

to allow maximal stem-

cell attachment. Cells (107/mL) were suspended in cell culture medium, and 5 L of cells

per scaffold piece were loaded into the polymer scaffold for a 5-min incubation period. For

the invitro studies, the cell-seeded PLG scaffolds were placed in wells of 12-well plates,

and the culture medium was changed every 23 days. At different time points, the cell-

seeded scaffolds were processed for standard histological and scanning electronic

microscopy (SEM) studies to observe their attachment and morphology. The SEM

procedures followed the protocol as described previously.10,15

Preparation of root fragments ofhuman teeth

Radicular portions of freshly extracted human teeth were horizontally sectioned into

segments of ~67 mm in length using an Isomet saw (Buhler, Lake Bluff, IL). The root

canal space was enlarged to ~12.5 mm in diameter. One end of the canal was sealed with

mineral trioxide aggregate (MTA) cement ~1 mm into the canal space, leaving the depth of

the canal space ~56 mm (Fig. 1). Root fragments were soaked at room temperature in 17%

ethylenediamine tetraacetic acid for 10 min and 19% citric acid for 1 min to remove the

smear layer,10,15

followed by treatment with betadine for 30 min and 5.25% NaOCl for 10

15 min for sterilization. Fragments were then rinsed in sterile PBS, soaked in PBS, and then

incubated at 37C for 37 days to remove residual sterilization agents and to ensure that

there is no microbial contamination.

FIG. 1. SCID mouse subcutaneous study model for pulp/dentin regeneration. The canal

space of human tooth root fragments (~67 mm long) was enlarged to ~12.5 mm indiameter. One end of the canal opening was sealed (more ...)


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FIG. 1.

SCID mouse subcutaneous study model for pulp/dentin regeneration. The canal space of human toothroot fragments (~67 mm long) was enlarged to ~12.5 mm in diameter. One end of the canal opening

was sealed with MTA cement. SCID, severe combined immunodeficient; MTA, mineral trioxideaggregate.

Insertion of cells/scaffolds into root fragments and implantation into mouse

subcutaneous space

Cells (107/mL) were seeded onto the PLG scaffolds for 5 min as described previously forin

vitro analysis. Immediately, the cells/PLG were inserted into the canal space of each root

fragment and kept in the wells of 12-well plates with a minimal amount of cell culture

medium. The tooth constructs were then implanted into the subcutaneous space on the back

of 6- to 8-week-old female severe combined immunodeficient mice (NOD.CB17-Prkdc-

scid/J; Jackson Laboratory, Bar Harbor, ME). Mice were anesthetized, and the dermal

space was created by blunt lateral dissection from a single dorsal midline incision. Each

mouse received two tooth fragments, one on each side. The wounds were sutured to obtain

primary closure. In parallel, root fragments with empty canal space were implanted into

severe combined immunodeficient mice as controls. Three to 4 months after the

transplantation, the mice were euthanized and the tooth fragments removed for histological

analysis. All animal procedures followed a protocol approved by the Institutional Animal

Care and Use Committee at the University of Maryland.

Histological examination

Samples (normal teeth and resected transplants) were fixed in 4% phosphate-buffered

paraformaldehyde, decalcified in 1015% ethylenediaminetetraacetic acid for 12 months,

paraffin embedded, longitudinally sectioned, and stained with hemotoxylin and eosin forhistological analysis. Some of the sections were used for immunohistochemical analysis.

Immunohistochemistry

Deparaffinized sections were immersed in 3% H2O2/methanol for 15 min to quench

endogenous peroxidase activity and incubated with primary antibodies (1:200 to 1:500

dilution) overnight at 4C. Primary antibodies used were as follows: alkaline phosphatase


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(ALP; LF-47), dentin sialoprotein (DSP, LF-21), and bone sialoprotein (BSP; LF-120),

provided by Dr. Larry Fisher (National Institute of Dental and Craniofacial Research,

National Institutes of Health, Bethesda, MD). Antibodies to CD105 were from BD

Biosciences and to human mitochondria were from Chemicon (Temecula, CA). Isotype-

matched control antibodies were used under the same conditions as the primary antibodies.

For enzymatic immunohistochemical staining, Zymed SuperPicTure polymer detection kit

(ZymedInvitrogen, Carlsbad, CA) was used according to the manufacturer's protocol.

Sections were counterstained with hematoxylin.


MethodsResultsDiscussionReferencesResultsIn vitro

charac

teriza

tion of den

tal s

tem cells

Cellmarkeranalysis

Human DPSCs and SCAP consisting heterogeneous populations of stem and progenitor

cells were reproducibly isolated using established protocols.811 Cells at passage 3 were

stained with antibodies against a number of cell surface markers and analyzed by flow

cytometry. CD14, CD34, and CD45 were not expressed, whereas CD73, CD90, and CD105

were expressed moderately to highly on these cells (Table 1). The surface marker

expression profile suggests that these cells were of typical mesenchymal cell lineages

similar to what normally is found in BMMSCs.16,17

Table 1. Surface Epitope Phenotype of Dental Stem/Progenitor Cells

a

MultipotentialityofSCAPandDPSCs

To verify that isolated dental cells qualified as stem/progenitor cells, we examined their

odonto/osteogenic, adipogenic, neurogenic, and myogenic potential with BMMSCs

included for comparison. Extracellular matrix production and mineralization by SCAP and

DPSCs after odonto/osteogenic stimulation was comparable to or greater than those of

BMMSCs (Fig. 2). SCAP and DPSCs were much weaker in adipogenicity because fewer

cells developed intracellular lipid droplet accumulation and showed delayed adipogenesis

compared with BMMSCs. Of those SCAP and DPSCs that did accumulate lipid droplets,


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they did not expand to the size of adipocyte-like cells that emerged from BMMSCs (Fig.

2B, E, H insets). Cells under neurogenic induction underwent morphological changes (not

shown). In general, BMMSCs exhibited spherical cell bodies with long and multiple

cellular extensions, whereas SCAP and DPSCs still possessed spindle-shaped fibroblastic

cell bodies with mainly long cellular processes. We found a subpopulation of these cells

(2030%) expressing the neurogenic marker III-tubulin (Fig. 2C, F, I). Under myogenic

stimulation, BMMSCs did not show any signs of myotube formation. SCAP and DPSCs

had a slender morphology and became elongated, but without myotube formation.

Myogenic gene expression analysis revealed that BMMSCs did not express any of the three

genes examined (myoD1, myogenin, and myosin heavy polypeptide 1), while myoD1 was

detected in SCAP and DPSCs after stimulation (Fig. 2J).

FIG. 2. Multiple lineage differentiation properties of SCAP and DPSCs compared with

BMMSCs. (A,D,G) Odonto/osteogenic induction and Alizarin Red staining ofmatrix mineralization. Insets: uninduced control. All scale bars including insets: 200

m. (more ...)

FIG. 2.Multiple lineage differentiation properties of SCAP and DPSCs compared with BMMSCs. (A,D,G)

Odonto/osteogenic induction and Alizarin Red staining of matrix mineralization. Insets: uninduced

control. All scale bars including insets: 200 m. (B, E,H) Adipogenic induction and Oil Red O staining

of the accumulated lipid droplets in cells. Inset images: higher magnification views of the adipocyte-likecells from the same culture shown in the main images. Scale bars: main images, 100 m; insets, 50 m.

(C,F,I) Neurogenic induction. III-tubulin: red fluorescence; 4',6-diamidino-2-phenylindole

dihydrochloride (DAPI): blue fluorescence. Scale bars: (C) 20 m; (F,I) 30 m. (J) Myogenic inductionand RT-polymerase chain reaction analysis of gene expression. M, RNA from cultured human myoblasts(obtained from Dr. Wesley M. Jackson, NIH/NIAMS, Bethesda, MD); C, uninduced control; m,

induction in myogenic medium. SCAP, stem cells from apical papilla; DPSC, dental pulp stem cells;BMMSCs, bone marrow mesenchymal stem cells.

Growth ofstemcellsinPLG

SCAP or DPSCs (5 105/50 L) were seeded onto a PLG copolymer scaffold (75:25 molar

ratio) cut into ~1.52 mm3, cultured invitro for 10 days or up to 8 weeks, and processed for

histological or SEM analysis. Cells seeded onto PLG attached well as revealed by SEM

analysis shown in Figure 3A, B, E, and F. The hemotoxylin and eosin staining showed that


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blank PLG scaffolds were completely colorless (not shown), suggesting that the eosin

staining seen in the DPSC-seeded PLG represented the cytoplasm of cells and extracellular

matrix (Fig. 3C, D). SEM analysis of the long-term cultures (8 weeks) revealed that cells

laid down fiber-like matrix (Fig. 3E, F). Minimal or no contraction of cells/PLG constructs

was noted after 78 weeks in cultures.

FIG. 3. Cell attachment and growth in the PLG scaffold invitro. Scanning electronic

microscopy of DPSCs seeded onto PLG after (A) 10 days and (B)14 days in culture.

(C,D) H&E staining of DPSCs seeded onto PLG after 7 weeks in culture. (E,F)Scanning (more ...)

FIG. 3.Cell attachment and growth in the PLG scaffold invitro. Scanning electronic microscopy of DPSCsseeded onto PLG after (A) 10 days and (B)14 days in culture. (C,D) H&E staining of DPSCs seeded

onto PLG after 7 weeks in culture. (E,F) Scanning electronic microscopy of SCAP seeded onto PLGafter 8 weeks in culture. Scale bars: (A) 200 m; (B,D, E) 20 m; (C) 250 m; and (F) 1 m. Arrows in(A) indicate attached cells, in (B) scaffold surface, in (E and F) fiber-like matrix. PLG, poly-D,L-lactide

and glycolide; H&E, hemotoxylin and eosin. Color images available online atwww.liebertonline.com/ten.

In vivo pulp/dentin tissue regeneration

Ingrowth ofsubcutaneoustissueintoemptiedrootcanalspace

The control root fragment did not receive any cell-seeded PLG in the emptied root canal

space. Three months after being transplanted into the mouse subcutaneous space, the canal

space appeared to be filled with the subcutaneous connective tissue, mainly adipose tissue,

which grew more than 2 mm into the canal (Fig. 4). The other half (to the MTA side) of the

canal space appeared empty, or the tissue was lost during the sample processing.

FIG. 4. Histological analysis of control group in the mouse subcutaneous model. (A) A root

fragment with empty canal space was transplanted into a SCID mouse for 3 months


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before being removed and processed for H&E staining. Mouse subcutaneous tissue

ingrown (more ...)

FIG. 4.Histological analysis of control group in the mouse subcutaneous model. (A) A root fragment with

empty canal space was transplanted into a SCID mouse for 3 months before being removed andprocessed for H&E staining. Mouse subcutaneous tissue ingrown from the canal opening (arrow). (B)

Higher magnification view showing the ingrown fatty tissue (arrow). Scale bars: (A) 1 mm; (B) 0.5 mm.

Vascularizedpulptissueregenerationinemptiedrootcanalspace

Longitudinal sections of the root constructs inserted with PLG seeded with SCAP or

DPSCs revealed that the emptied canal was filled with regenerated pulp-like tissue as

shown in Figures 5 and and6.6. Voids were scattered in the pulp-like tissue, which was

most likely the result of unresorbed PLG scaffolds. There was a sharp distinction between

the histological characteristics of the regenerated pulp-like tissue situated only within the

canal space and those of the subcutaneous soft tissues in the mouse dermal space. A thin

layer of fibrous capsule separated the two types of tissues. The entire pulp-like tissue was

vascularized with a uniform cell density resembling the natural pulp (Fig. 5I).

FIG. 5. Histological analysis ofinvivo pulp/dentin regeneration using SCAP. A root

fragment was prepared and the canal space inserted with SCAP/PLG andtransplanted into a SCID mouse for 3 months. The sample was harvested and

processed for H&E staining. (more ...)

FIG. 5.Histological analysis ofinvivo pulp/dentin regeneration using SCAP. A root fragment was prepared and

the canal space inserted with SCAP/PLG and transplanted into a SCID mouse for 3 months. The samplewas harvested and processed for H&E staining. D, original dentin; rD, regenerated dentin-like tissue; rP,

regenerated pulp-like tissue. Blue arrow in (A) indicates the blood supply entrance; green arrows in (B)

and (C) indicate continuous layer of uniformed thickness of rD; yellow arrows in ( E) and (F) indicatethe region of well-aligned odontoblast-like cells with polarized cell bodies; green arrows in (G and H)


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indicate junctions between D and rD. Scale bars: (A) 1 mm; (B and C) 500 m; (D) 100 m; (E and F)

20 m; and (GI) 50 m.

FIG. 6. Histological analysis ofinvivo pulp/dentin regeneration using DPSCs. Samples were

prepared using the same procedures as described in Figure 5, except that the samplewas harvested from the SCID mouse 4 months after implantation. D, original dentin;

(more ...)

FIG. 6.Histological analysis ofinvivo pulp/dentin regeneration using DPSCs. Samples were prepared using the

same procedures as described in Figure 5, except that the sample was harvested from the SCID mouse 4

months after implantation. D, original dentin; rD, regenerated dentin-like tissue; rP, regenerated pulp-like tissue. Od, odontoblast-like cells. Green arrows in (A) indicate rD; blue arrows in (A) indicate the

entrance of blood supply; blue arrows in (B and C) indicate the thin layer of rD under MTA cement;

blue arrows in (F and G) indicate the junction of D and rD; black arrow in (G) indicates well-aligned

odontoblast-like cells. Yellow arrows in (G) indicate dentinal tubule-like structures. Scale bars: (A) 1mm; (B) 200m; (CE) 100 m; and (F and G) 50 m. Color images available online atwww.liebertonline.com/ten.

Formationofdentin-liketissueandodontoblast-likecells.

A representative sample using SCAP/PLG showed a continuous layer of mineralized tissue

with uniform thickness (~200 m) deposited onto the existing dentinal walls and onto the

MTA cement surface in the canal (Fig. 5); herein referred to as regenerated dentin-like

tissue. When using DPSCs/PLG, a layer of regenerated dentin-like tissue was also

deposited onto the canal dentinal walls (Fig. 6). In this sample, the regenerated dentin-like

tissue had less continuity and thickness compared with the sample shown in Figure 5.

Higher magnification observations revealed that a layer of odontoblast-like cells were linedagainst the mineralized dentin-like tissue (Figs. 5 and and6).6). There were also scattered

cells embedded within the dentin-like structure. Some located between the original dentin

and the regenerated dentin. The newly deposited dentin-like tissue appeared to tightly

adhere to the original dentin except where there were gaps that resulted from sample

processing artifacts. The newly formed mineralized tissue also appeared to fill into the


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space of dentinal tubules (Fig. 5H). At higher magnification, unlike the natural dentin, the

pulp side of the regenerated dentin-like tissue was not smooth; instead, there were

projections of the structures into the pulp side (Figs. 5 and and66).

The odontoblast-like cells were not well organized, nor well aligned as the natural

counterparts, and it was difficult to observe the typical characteristics that natural

odontoblasts possess (e.g., polarized cell bodies). Nonetheless, some regions of the

odontoblast-like cells showed somewhat typical odontoblast characteristics. Groups of

odontoblast-like cells were well aligned, such that each cell was spatially juxtaposed to

adjacent cells with overt polarized morphology (Fig. 5E, F, and Fig. 6G). The regenerated

dentin-like tissue did not form observable well-organized dentinal tubules, except in a few

regions where odontoblast-like cells were better aligned (Fig. 6G).

Cellsinregeneratedpulp-liketissueareofhumanorigin.

To verify that the cells responsible for generating the pulp/dentin-like tissues in the canal

space were of human origin, antibodies against human mitochondria were used to detect

human cells in the tissues. As shown in Figure 7DF, a majority of the cells stained

positively for the antibodies, confirming the human cell origin of these cells in the

regenerated pulp-like tissue.

FIG. 7. Immunohistochemical analysis of pulp tissue regenerated invivo using SCAP. (AC)

H&E staining; immunostaining for (DF) human mitochondria; (G) dentinsialoprotein; (H) alkaline phosphatase and (I) bone sialoprotein. D, original (more ...)

FIG. 7.Immunohistochemical analysis of pulp tissue regenerated invivo using SCAP. (AC) H&E staining;

immunostaining for (DF) human mitochondria; (G) dentin sialoprotein; (H) alkaline phosphatase and

(I) bone sialoprotein. D, original dentin; rD, regenerated dentin-like tissue; rP, regenerated pulp-liketissue. Arrows indicate cells with positive staining. Color images available online atwww.liebertonline.com/ten.


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Odontoblast-likecellsexpressodontogenicmarkers

To further verify the dentin-like tissue was generated by cells of odontoblast lineage,

several odonto/osteogenic markers were examined immunohistochemically. Positive

staining for DSP, BSP, and ALP was seen in cells lining against as well as those embedded

in the newly deposited dentin-like tissues, suggesting that the differentiated odontoblast-

like cells were responsible for the production of the calcified tissue (Fig. 7GI).

In addition, we compared CD105 expression in the regenerated and the natural pulp.

CD105 was expressed by DPSCs and SCAP based on the flow cytometry analysis

presented in Table 1. Immunocytochemical analysis of SCAP in cultures confirmed their

CD105 expression (Fig. 8A). In the natural pulp, CD105 expression was detected in

association with blood vessels and in odontoblasts (Fig. 8C, D). CD105 (Endoglin) is

known to be expressed on human vascular endothelial cells. In comparison, CD105 was

also expressed by odontoblast-like cells and paravascular cells in the regenerated pulp-like

tissus (Fig. 8E, F), suggesting that the regenerated tissue closely resembles natural pulp.

FIG. 8. Immunocyto/histochemical staining of CD105. Cultured SCAP immunostained with

(A) anti-CD105 antibody or (B) nonimmune IgG as negative control. (C) Positivestaining of odontoblasts (arrowheads) in a sample of natural human dental

pulp/dentin tissue. ( (more ...)

FIG. 8.Immunocyto/histochemical staining of CD105. Cultured SCAP immunostained with (A) anti-CD105

antibody or (B) nonimmune IgG as negative control. (C) Positive staining of odontoblasts (arrowheads)in a sample of natural human dental pulp/dentin tissue. (D) A blood vessel in the natural pulp is positive

for CD105 (arrow). (E) Positive immunostaining is seen in odontoblast-like cells lining against rD or

inside rD (arrowheads). (F) CD105-positive cells are seen in rP and around a blood vessel (arrow). rD,

regenerated dentin-like tissue; rP, regenerated pulp-like tissue. Color images available online atwww.liebertonline.com/ten.


MethodsResultsDiscussionReferencesDiscussionOur study is the first invivo evidence demonstrating denovo synthesis of vascularized

human pulp/dentin-like tissues in an emptied human root canal space 56 mm deep with an


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opening end of only ~2.5 mm. The most striking phenomenon is the formation of a

continuous layer of dentin-like tissue on the existing canal dentinal walls and on MTA

cement surface. Expression analysis of several key genes, including DSP, BSP, ALP, and

CD105, indicates that the regenerated pulp tissues closely resemble natural pulp tissue. Our

findings constitute the first step toward stem cellmediated denovo regeneration of

pulp/dentin in clinical endodontic practice. Use of heterogeneous population of

stem/progenitor cells appeared to be able to achieve this purpose. Both SCAP and DPSCs

as mesenchymal-like stem cells presented a different profile of multipotency from

BMMSCs. This may be because of their different developmental originderived from or

associated with neural crest cells.

Previously, we discussed several issues regarding the establishment of a protocol for

pulp/dentin regeneration, including the scaffolds suitable for this purpose, the extent of

vascularization of the regenerated pulp, and the formation of odontoblasts and new dentin

on the existing dentin surface.4,5,10,16

Our previous invitro studies showed that although

collagen gels are a convenient cell carrier for pulp regeneration in the root canal space,

severe contraction may hamper the regeneration process.10

A recent invivo study targeting

root dentin perforation repair with the use of DPSCs and dentin matrix protein 1 carried by

collagen matrix showed that soft connective pulp-like tissue was formed in the perforated

site, but no hard tissue was generated.18

Regeneration of a small portion of pulp after

pulpotomy using a collagen matrix appears to be successful as demonstrated in a dog study

model.19

Biosynthetic scaffolds such as PLG or poly-L-lactic acid are much more resistant

to contraction than collagen. They are generally made into porous form to provide inner

spaces for cells to populate. Cordeiro etal. reported successful regeneration of well-

vascularized pulp-like soft tissue utilizing poly-L-lactic acid as a scaffold fabricated in the

pulp chamber space of a tooth slice that was seeded with stem cells from human exfoliated

deciduous teeth.6Together with our findings in this study, synthetic polymers of lactide

and/or glacolide appeared to be suitable scaffold systems fordenovo pulp regeneration. We

speculated that the slow degradation of the PLG in our study negatively affected the quality

of the regenerated pulp-like tissue. In future studies, modification of the ratio of lactide and

glycolide or selection of other type of scaffold materials such as silk may improve this

situation.20,21


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Vascularization in the regenerated pulp tissue in canal space was a major concern because

the blood supply for the space can only come from the apex.4The above mentioned studies

have tried to avoid this issue in testing pulp regeneration by using a thin tooth slice

model.6,18 Utilizing factors such as vascular endothelial growth factor and/or platelet-

derived growth factor to enhance angiogenesis or the addition of vascular cells or

hematopoietic progenitors has been an effective approach to resolve to this issue for tissue

regeneration.14,2225

In this study, angiogenic enhancement was not employed. The tooth

fragments contained a canal space 56 mm deep; therefore, it was likely that the blood

vessel growth into the end of the canal space to provide nutrients for the stem cells could

not occur shortly after the tooth fragment was transplanted into the mouse subcutaneous

space. However, our results indicate that cells not only survived well but also were able to

regenerate tissues. It is possible that the nutrients were able to diffuse into the canal space,

allowing the stem cells to build new tissues; once the blood vessels were generated in the

entire canal space, long-term stability of the tissue was established. The question arises as

to whether a smaller opening of the canal, that is,


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dentin did not appear to affect this capacity. From these observations, it appears that stem

cells seeded in the scaffold are attracted to the dentinal wall, differentiate into odontoblast-

like cells, and extend their cellular processes into the dentinal tubules. The mechanism

underlying this phenomenon has been speculated to be the release of growth factors such as

transforming growth factor- that is known to be embedded in dentin,26,27

which attracts

and induces odontoblast differentiation of the seeded stem cells.10

Chemical disinfection of

the root canal space may damage these embedded growth factors. Further investigation is

needed to seek ways to avoid this potential damage.

The most interesting and important finding of this study is the formation of a continuous

layer of dentin-like tissue with uniform thickness on the existing canal dentin walls and the

MTA cement surface, especially with SCAP-seeded samples. This fulfills, at least in part, a

major requirement of functional tissue engineering/regeneration because dentin production

is one of the major functions of pulp. However, the characteristics of the regenerated

dentin-like tissue are quite different from the natural dentin. First, dentinal tubules are not

obvious, possibly because of their convoluted paths. Second, it is quite cellular, and third,

the formation is not synchronous. There were only some regions of the new dentin-like

tissue that showed dentinal tubule-like structures (Fig. 6G). Our previous invitro work

demonstrated that DPSCs seeded onto a processed dentin surface morphologically

transformed into odontoblast-like cells each with a cellular process extending into the

dentinal tubule.10,15

Based on this observation along with the present invivo findings, the

following events were likely to have occurred subsequent to invivo transplantation of the

tooth fragments: (i) some SCAP or DPSCs on PLG in the canal space migrate toward the

dentin surface, (ii) these cells receive signals from dentin and differentiate into odontoblast-

like cells, (iii) a cellular process is extended from each cell into the dentinal tubules, (iv)

cells initiate production of extracellular matrix in the dentinal tubule space as well as onto

the dentin surface, and (v) the remaining steps of dentin-like tissue apposition are similar to

the natural dentin production, except cells laid down dentin-like tissue at a different pace.

The question remains as to whether every dentinal tubule is occupied by a newly

differentiated odontoblast-like cell of equal potentiality. If not, it may explain the

disorganized nature of the newly synthesized dentin-like tissue and the alignment of those

odnotoblast-like cells as well as the entrapment of many of these cells in the dentin-like


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structure. Tertiary dentin produced by the natural pulp has less organization of dentinal

tubules and may have cell entrapment in the dentin; therefore, the newly regenerated

dentin-like tissue may be similar to tertiary dentin.

Taken together, our findings show that pulp/dentin regeneration can be accomplished with

a stem/progenitor cellmediated tissue engineering approach. This study is the first step

toward reaching the goal of regenerating functional pulp/dentin that is very similar to its

natural counterparts. To achieve this goal, future investigations must include the use of

larger animals, such as minipigs, to regenerate these tissues insitu in the jaw bone. Besides

gaining vascularity, the innervation of the regenerated pulp/dentin tissue by ingrowth of

nerve fibers from the apical tissues requires extensive investigation. Further, if a higher

quality of the regenerated dentin tissue is desired, that is, producing new dentin with well-

organized dentinal tubules, the method of treating dentin surface may need modification to

provide better signaling stimulation to guide stem cell differentiation toward odontoblast

lineage in a more synchronous manner.

AcknowledgmentsThis work was supported in part by grants from the National Institutes of Health (NIH) R01

DE019156-01 (G.T.-J.H.), RO1 DE17449 (S.S.), by the NIAMS/NIH Intramural Research

Program Z01 AR41131 (R.S.T), and by an Endodontic Research Grant from American

Association of Endodontists Foundation (G.T.-J.H.). We wish to thank Dr. MatthewDaniels (NIH/NHLBI, Electron Microscopy Core) for the assistance of SEM analysis, Dr.

Larry Fisher (NIH/NIDCR) for providing antibodies, Dr. Wesley M. Jackson

(NIH/NIAMS) for providing the RNA from cultured human myoblasts, and Dr. Yi Liu

(USC) for the assistance of neurogenic studies.

Disclosure StatementNo competing financial interests exist.


MethodsResultsDiscussionReferencesReferences

1. Mooney D.J. Powell C. Piana J. Rutherford B. Engineering dental pulp-like tissue in vitro. BiotechnolProg.

1996;12:865. [PubMed]


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2. Bohl K.S. Shon J. Rutherford B. Mooney D.J. Role of synthetic extracelluar matrix in development of engineered

dental pulp. JBiomaterials Science.(Polymer Edition) 1998;9:749.

3. Buurma B. Gu K. Rutherford R.B. Transplantation of human pulpal and gingival fibroblasts attached to synthetic

scaffolds. EurJOralSci.1999;107:282. [PubMed]

4. HuangG

.T

.J. Sonoyama W. Liu Y. Liu H. Wang S. Shi S.T

he hidden treasure in apical papilla: the potential rolein pulp/dentin regeneration and bioroot engineering. JEndod.2008;34:645. [PMC free article] [PubMed]

5. Huang G.T.-J. Apexification: the beginning of its end. IntEndodJ.2009;42:855. [PubMed]

6. Cordeiro M.M. Dong Z. Kaneko T. Zhang Z. Miyazawa M. Shi S. Smith A.J. Nor J.E. Dental pulp tissue

engineering with stem cells from exfoliated deciduous teeth. JEndod.2008;34:962. [PubMed]

7. Batouli S. Miura M. Brahim J. Tsutsui T.W. Fisher L.W. Gronthos S. Robey P.G. Shi S. Comparison of stem-

cell-mediated osteogenesis and dentinogenesis. JDentRes.2003;82:976. [PubMed]

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