Human Reproduction vol.13 no.10 pp.2857–2868, 1998
Mitochondrial transfer between oocytes: potential
applications of mitochondrial donation and the issue of
heteroplasmy
Jonathan Van Blerkom1, Jane Sinclair and
Patrick Davis
Department of Molecular, Cellular, and Developmental Biology,
University of Colorado, Boulder, CO 80309, USA
1To
whom correspondence should be addressed
The developmental competence of mouse and human early
embryos appears to be directly related to the metabolic
capacity of a finite complement of maternally inherited
mitochondria that appear to begin to replicate after
implantation. Mitochondrial dysfunctions resulting from a
variety of intrinsic and extrinsic influences, including
genetic abnormalities, hypoxia and oxidative stress, can
profoundly influence the level of ATP generation in oocytes
and early embryos, which in turn may result in aberrant
chromosomal segregation or developmental arrest. Deletions and mutations in oocyte mitochondrial DNA may
subtend metabolic deficiencies or replication disorders in
some infertile women and in women of increased reproductive age. Here, we describe methods for (i) the compartmentalization of mouse and human oocyte mitochondria
into unique cytoplasts enriched for these organelles, and
(ii) their transfer by microinjection into intact recipient
oocytes. Metabolically active mitochondria in donor and
recipient metaphase II stage oocytes were labelled with
mitochondria-specific fluorescent probes, and the fate and
location of donated mitochondria in recipient oocytes were
followed by conventional epifluorescence and scanning laser
confocal fluorescence microscopy. The net ATP content of
undisturbed and recipient oocytes from the same cohort(s)
was measured quantitatively at timed intervals after mitochondrial injection. The results demonstrate the feasibility
of isolating and transferring mitochondria between oocytes,
an apparent increase in net ATP production in the recipients, and the persistence of activity in the transferred
mitochondria. The findings are discussed with respect to
mitochondrial function and dysfunction in mammalian
oocytes and embryos, and to the potential clinical applications of mitochondrial donation as they relate to the creation
of heteroplasmic embryos.
Key words: heteroplasmy/mitochondria/mitochondrial donation/mitochondrial function in pre-implantation embryos
Introduction
Mitochondria are self-replicating, maternally inherited
organelles that utilize oxidative phosphorylation pathways to
supply ATP for all energy-requiring cellular activities. These
© European Society for Human Reproduction and Embryology
organelles are the primary genetic elements which contribute
to patterns of cytoplasmic inheritance that can profoundly
effect development, metabolism, and reproductive performance
(Smith and Alcivar, 1993). In mature (metaphase II, MII)
mammalian oocytes and early cleavage stage embryos, mitochondria are structurally undifferentiated and generate ATP at
relatively low levels when compared to those observed at the
late morula and blastocyst stages (Van Blerkom, 1989, for
review). In the human, significant differences in net ATP
content occur among mature oocytes from the same and
different patients, and these differences are not only oocytespecific but appear to be associated with embryo developmental
competence (Van Blerkom et al., 1995a; Barnett and Bavister,
1996). In this respect, several investigators have suggested that
the maternal age-associated reduction in embryo developmental
competence may be related to an inadequate capacity to
generate ATP at levels sufficient to support normal chromosomal segregation (Gaulden, 1992) or normal biosynthetic,
mitotic, and physiological activities within blastomeres, with
developmental abnormalities or arrest during the preimplantation stages being the result (Van Blerkom et al.,
1995a; Barnett and Bavister, 1996). Mitochondrial dysfunction
leading to oxidative damage and apoptosis, hypoxia, and
deletions or point mutations in the oocyte mitochondrial
genome (mtDNA) (Corbisier and Remacle, 1990, 1993,
Clayton, 1992; Wallace, 1992; Shigenaga et al., 1994;
Lightowlers et al., 1997), especially in oocytes of older
women (Keefe et al., 1995), are the types of adverse influences
that may contribute to reduced mitochondrial function in
the human female gamete. Because significant mitochondrial
replication may not occur until the hatched blastocyst stage
(see Van Blerkom, 1989; Smith and Alcivar, 1993, for reviews),
or in some species such as the mouse until the egg cylinder
stage (Ebert et al., 1988) all energy-requiring activities for the
embryo are largely dependent upon the normal function of
a finite mitochondrial complement (~100 000 mitochondria/
oocyte, Chen et al., 1995) that is partitioned with each cell
division among the blastomeres of cleaving embryos and
the inner cell mass and trophoblast cells of the developing
blastocyst. Consequently, any potential adverse influence(s) on
normal mitochondrial function and differentiation during the
preimplantation stages, even if only a portion of the mitochondria are affected, could be of direct developmental significance for the embryo.
If mitochondrial dysfunction or reduced metabolic capacity
contribute to the developmental incompetence of preimplantation embryos, then efforts to increase metabolism
constitutively rather than transiently may be beneficial if
embryo demise or developmental abnormality are likely out2857
J.Van Blerkom, J.Sinclair and P.Davis
comes. Transfer to early mouse embryos of genetically marked
mouse and hamster mitochondria isolated from somatic cells
(e.g. testis, liver) has no apparent effect on embryogenesis as
indicated by normal gestation and term births. However, the
donated mitochondria either fail to replicate or do not persist
in quantities detectable by molecular analysis (Ebert et al.,
1989). In this respect, the introduction of presumably normal
and functional mitochondria derived from a competent oocyte
may enhance metabolic activity in oocytes where mitochondrial
defects or metabolic deficiencies are suspected. In addition,
owing to the origin and relatively undifferentiated state of
oocyte mitochondria, the ‘donor’ mitochondria may have a
higher probability of integrating with the endogenous population and be subject to the same processes that regulate their
differentiation, with an attendant increase in the probability of
their replication, entrance into the germ line, and persistence
in the adult.
The purpose of the present study was to determine whether
mitochondria could be harvested from one oocyte and introduced into another. The results demonstrate that mouse and
human oocyte mitochondria can be segregated by centrifugation
into a cytoplasmic compartment or cytoplast that is free of
MII chromosomes, aspirated into a micropipette and, for the
mouse, inserted into a mature oocyte. The donor mitochondria
progressively migrate from the site of insertion and can be
identified in the living recipient mouse oocytes for at least
80 h. The results suggest that mitochondrial donation from
one oocyte to another is possible, and the findings are discussed
with respect to (i) mitochondrial function and dysfunction in
oocytes and preimplantation embryos, (ii) potential clinical
applications in the treatment of infertility and inherited mitochondrial disorders, and (iii) issues related to heteroplasmy.
Materials and methods
Oocyte collection, culture, fluorescent probe loading and
centrifugation
MII stage oocytes were obtained from the oviducts of sexually mature
mice at 12–14 h after the administration of a superovulatory dose of
human chorionic gonadotrophin (HCG) (5 IU) to animals primed
with pregnant mare serum gonadotrophin (5 IU) 48 h earlier. Germinal
vesicle stage oocytes (GV) were harvested from small antral follicles,
denuded of cumulus and coronal cells by repeated passage through
a glass micropipette and cultured in medium M2 to MII as previously
described (Van Blerkom et al., 1995a). Prior to centrifugation, the
zona pellucida was removed by exposure to warm (37°C) acidic
Tyrode’s solution for 10–15 s. To visualize mitochondria in living
oocytes, zona-free oocytes were cultured in M2 supplemented with
one of the following mitochondria-specific fluorescent probes: (i)
rhodamine 123 (10 µg/ml, 4 min), (ii) Mitofluor Green (0.5 µg/ml,
30 min) or (iii) Mitotracker Red or Mitotracker Green (0.5 µg/ml,
30 min). Rhodamine 123 and Mitotracker Red only label mitochondria
that are metabolically active. For some experiments, staining with
the DNA-specific probe 49,6-diamidino-2-phenylindole diacetate
(DAPI, 5 µg/ml, 5 min) was performed in order to detect mitochondrial
DNA and chromosomal fluorescence in living oocytes. Chromosomal
fluorescence was detected in glutaraldehyde- or formaldehyde-fixed
specimens with YOPRO-1 iodide (5 µg/ml, 6 min). All fluorescent
probes were obtained from Molecular Probes (Eugene, OR, USA)
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and protocols for staining followed procedures previously described
(Van Blerkom et al., 1995b).
After preloading of the fluorescent probes, zona-free oocytes were
cultured in the presence of cytochalasin D (CCD, 1 µg/ml, 1 h) and
then transferred in 1.0 ml of medium M2 containing 1.0 µg/ml CCD
to the surface of a 48% Percoll solution (1 ml) (Sigma, St Louis,
MO, USA) prepared in HEPES-buffed M2. Oocytes were centrifuged
at 19 600 g for 2 h and specimens were recovered from the lower
portions of the centrifuge tube. Mitochondria-rich and mitochondriapoor cytoplasts were identified by morphology and fluorescence and
transferred to 20 µl microdroplets of M2. Representative samples
were fixed for fine structural analysis in protein-free M2 containing
1% glutaraldehyde and prepared for transmission electron microscopy
(Van Blerkom et al., 1995b).
Normal-appearing, uninseminated MII-stage human oocytes that
were in excess of the number replaced in the Fallopian tube in gamete
intra-Fallopian transfer (GIFT) procedures were denuded of cumulus
and coronal cells, preloaded with mitochondria-specific fluorescent
probes, and cultured in the presence of CCD (10 µg/ml) for 1.5 h.
After removal of the zona pellucida, oocytes were centrifuged for
2.5 h as described above.
Mitochondrial transfer by microinjection
Recipient mouse oocytes that had matured in vivo or in vitro were
incubated in M2 supplemented with CCD (1 µg/ml) for 1–2 h prior
to injection. For microinjection, oocytes were transferred to a 20 µl
microdroplet of M2 under mineral oil in a modified 35 mm tissue
culture dish. The temperature of the microdroplet was maintained at
precisely 37°C by means of an electric current passed through a
thermo-optically treated glass coverslip integrated into the bottom of
the plastic culture dish in a ∆T Culture Dish System (Bioptechs,
Butler, PA, USA) as previously described (Van Blerkom et al., 1995b).
Micropipettes were prepared with an internal diameter of ~5–7 µm
for injection and 10–15 µm for holding. Injection pipettes were
bevelled to an angle of 40° and both injection and holding pipettes
were bent to an angle of 30–40° from horizontal in a microforge.
Mitochondria-enriched cytoplasts were held under negative pressure
and a portion of the contents was aspirated manually into the injection
pipette. Lysis of the cytoplasts during aspiration was relatively rare,
occurring in ,10% of these structures. In some instances, multiple
aspirates were obtained from the same portion of the cytoplast that
appeared to contain the highest concentration of mitochondria. The
cytoplast was removed and replaced with an intact (zona-enclosed)
oocyte held with the first polar body at the 12 o’clock position. A
previously determined aspirate volume between 5 and 10 pl was
expressed by manually adjusting the rate of delivery (Nikon PLI-188
Microinjection System). The injection pipette was displayed on a
high resolution video monitor at 3200 magnification and cytoplasm
was aspirated into the pipette until it reached a mark on the video
screen corresponding to predetermined volumes between 5 and
10 pl. Insertion into the oocyte usually occurred at the 3 o’clock
position with deposition either in the approximate centre of the oocyte
or in a pericortical location. Other types of deposition included
placement at multiple locations and a progressively increasing deposition of mitochondria as the pipette was withdrawn from the centre
to the periphery of the oocyte. Immediately after injection oocytes
were fixed or returned to normal culture medium. At 12 h intervals
up to 80 h after injection, at which time oocyte culture was terminated,
recipient oocytes were either fixed or examined in the living state.
Detection of mitochondrial and DNA-specific signals in living and
fixed cytoplasts and recipient oocytes utilized conventional and
scanning laser confocal fluorescence microscopy as previously
Interoocyte mitochondrial transfer
described (Van Blerkom et al., 1995b). Mitochondrial transfers
between human oocytes were not undertaken in this study.
Measurements of ATP content in cytoplasmic aspirates, intact and
injected mouse oocytes
The ATP content of (i) cytoplasts, (ii) karyoplasts and (iii) untreated
and recipient oocytes from the same cohort(s) was determined
quantitatively by measurement of luminescence generated in an ATPdependent luciferin–luciferase bioluminescence assay as previously
described (Van Blerkom et al., 1995a). The same volume of cytoplasmic aspirate injected into an oocyte was expressed into distilled
water and prepared for ATP determination with the same protocol
used for intact oocytes. A standard curve containing 17 ATP concentrations from 5 fmol to 5 pmol was generated for each series of analyses.
Results
Stage-specific effects of centrifugation on mouse oocyte
morphology and intracellular organization
The efficacy of different iso-osmotic media (sucrose,
Nycodenz, Percoll), gradient types (step, continuous,
and single concentration), centrifugation times and forces,
and agents which depolymerize microtubules (nocodozole) and
microfilaments (cytochalasin B and D), either individually or
in combination, was examined in preliminary studies. These
studies led to a standard experimental protocol which used a
single concentration of Percoll, preincubation of mouse oocytes
in CCD for 1 h, and centrifugation for 2 h. In-vitro matured,
CCD-treated oocytes were centrifuged at the GV, germinal
vesicle breakdown (GVBD), circular bivalent (CBV) and
metaphase I (MI) and MII stages. Oocytes were exposed to
CCD because preliminary studies showed that no significant
alteration in oocyte geometry or cytoplasmic organization
occurred in untreated oocytes. Prior to MI, only a slight
distortion from the normal spherical shape was observed in
CCD-exposed oocytes, which appeared slightly elongated or
oval after centrifugation. For example, at the light microscope
level the sole indication that centrifugation affected the GV
stage oocyte was a distortion of the nuclear membrane from
a spherical structure (Figure 1A) to one which appeared bellshaped (Figure 1B, C). However, even these changes in
nuclear geometry were dependent upon the time following the
resumption of meiosis. Between 0 and 10 min of culture,
centrifugation had a minimal influence on nuclear shape,
although a tendency towards an oval appearance was observed
(Figure 1A). A significant change in nuclear geometry was
first evident at ~30 min (Figure 1B) to 40 min of culture
(Figure 1C), with the appearance of a bell-shaped germinal
vesicle in which the nucleolus was oval to slightly elongated
(Figure 1B, C). Figure 1D indicates the distribution of DNA
staining in an elongated GV. A very similar situation prevailed
at the GVBD and circular bivalent (CBV) stages where
chromosomes were displaced to one pole of an otherwise
spherical oocyte (arrow, Figure 1E).
The first clearly detectable effect of centrifugation on oocyte
shape occurred just prior to or at MII. MII-stage oocytes
(in-vivo or in-vitro matured) elongated progressively and
significantly during centrifugation (Figure 1F, G, H) and
the cytoplasm appeared to segregate into a largely granular
compartment (black asterisk, Figure 1F) and a largely translucent compartment (white asterisk, Figure 1F). Two characteristic shapes were observed after ~60 min of centrifugation,
a ‘dumbbell-like’ structure such as the one shown in Figure
1F, and an elongated ‘tube-like’ structure with a bulbous end
such as the one shown in Figure 1H. After ~90 min of
centrifugation, these compartments separated into granular and
agranular cytoplasts (Figure 1N, P, R) with the first polar body,
if still present, associated with the agranular structure (Figure
1I). When placed in normal culture medium, agranular cytoplasts assumed a spherical shape (Figure 1R, T, U) while the
granular compartments became pear-shaped (Figure 1R).
Fine structural analysis of centrifuged GV stage oocytes at
timed intervals after the resumption of meiosis confirmed the
light microscopic finding that the GV was relatively resistant
to distortion in the uncultured oocyte (t 5 0 min, Figure 2A)
but elongated progressively with culture (t 5 45 min, Figure
2B) and the approach of GVBD (t 5 60 min, Figure 2C). In
contrast, a displacement of cytoplasmic organelles in the
direction of the maximal centrifugal force (large arrows, Figure
2A–C) was detected in uncultured GV-stage oocytes (asterisk,
Figure 2A), and the magnitude of this displacement increased
significantly as maturation progressed to GVBD (asterisk,
Figure 2B–C). Compartmentalization of organelles and other
cytoplasmic structures in GV stage oocytes was detected during
the GV stage but was particularly evident when oocytes
were matured for ~45 min prior to centrifugation where
mitochondria and cytoplasmic lattices appeared to be
migrating to one hemisphere of the oocyte (asterisk, Figure
2C). Zones of cytoplasm devoid of mitochondria (insert, Figure
2A) and cytoplasmic lattices are indicated by an asterisk in
Figure 2A–D, and in the same figures, the apparent boundary
between organelle-rich and organelle-poor regions of the cytoplasm is denoted by arrowheads. At the GVBD/CBV stage,
the portion of centrifuged oocytes depleted of mitochondria
contained small coated vesicular structures, membranous
elements, lipid bodies and a low density of cytoplasmic lattices
(arrows, Figure 2G).
Generation of mitochondria-enriched cytoplasts and mitochondria-deficient karyoplasts in metaphase II stage oocytes
Mouse
The two types of structures generated by the centrifugation of
MII oocytes differed in size, granularity, and cytoplasmic
contents. Light (Figure 1K, M, P, R) and fluorescent microscopic analysis with mitochondria- (Figure 1G, M, O, Q, S)
and DNA-specific probes (Figure 1G, H, J, T, U) demonstrated
that the agranular, smaller structures were karyoplasts that
contained chromosomes, a single cluster of lipid bodies (arrow,
Figure 1K, asterisk Figure 1L) and virtually no detectable
mitochondrial fluorescence. In Figure 1R, karyoplasts are
indicated by a white asterisk and cytoplasts by a black
asterisk. The same specimens analysed for mitochondrial
fluorescence are shown in Figure 1S, where the white asterisks
indicate the positions of the karyoplasts, in which chromosomal
fluorescence was confirmed by DAPI staining (arrows, Figure
1T–U). The segregation of the cytoplasm into mitochondria
deficient and mitochondria enriched compartments was evident
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J.Van Blerkom, J.Sinclair and P.Davis
prior to the separation of the oocyte into two distinct types of
‘cytoplasts’. Figures 1K and M are representative examples of
an MII oocyte subjected to centrifugation for ~45 min in
which a significantly higher intensity of mitochondrial fluorescence was observed in one compartment (asterisk, Figure
1M). After 70 min of centrifugation, MII oocytes assumed a
2860
characteristic ‘dumb-bell-like’ shape (Figure 1F) in which the
segregation of mitochondria and chromosomes into separate
compartments appeared to be virtually complete, as indicated
by the presence or absence of a respective fluorescent signal
(Figure 1G). A very similar distribution was detected in oocytes
that assumed a tubule-like shape in which all of the detectable
Interoocyte mitochondrial transfer
chromosomal fluorescence was localized to one end of the
oocyte (arrow, Figure 1H) and a cluster of DAPI-stained
mitochondria was localized at the other end (asterisk, Figure
1H).
The separation of centrifuged MII oocytes into mitochondriaenriched cytoplasts and mitochondria-deficient karyoplasts was
confirmed by fluorescent probe analysis and transmission
electron microscopy. Figures 3A and B show the results of a
typical separation in which MII oocytes were stained with
DAPI prior to centrifugation. The two classes of cytoplasts
evident with differential interference contrast optics differed
in size, cytoplasmic granularity (asterisks, Figure 3A) and
intensity of DNA staining (Figure 3B). The presence of
mitochondria in the cytoplasts with higher levels of DAPI
fluorescence (black asterisks, Figure 3B) was demonstrated
with mitochondria-specific probes (black asterisks, Figure 1Q,
S). At higher magnifications, MII chromosomes in separated
oocyte compartments with low DAPI fluorescence (white
asterisks, Figure 3B) were localized to one pole of the
nucleoplast (Figure 1T, U) in which little, if any, mitochondrial
fluorescence was detected (e.g. Figure 1S). Serial optical
sections taken by scanning laser confocal microscopy revealed
that mitochondria were often clustered at one pole of the
cytoplast (asterisk, Figure 3C). This mitochondria-dense zone
could be identified with differential interference contrast optics
owing to the increased granularity associated with this
region (asterisk, Figure 3A), and its detection was used to
orient the cytoplast on the holding pipette in order to maximize
the retrieval of mitochondria (see below).
Fine structural analysis demonstrated the scarcity and
abundance of mitochondria in the karyoplasts (Figure 2E–F)
and cytoplasts (Figure 2I) respectively. In some sections of
karyoplasts, assemblages of Golgi saccules (Figure 2E) and
distended cisternae of the smooth-surfaced endoplasmic
reticulum (Figure 2H) were detected in the cortical cytoplasm.
However, as shown in Figure 2E, the cytoplasm of karyoplasts
was generally devoid of organelles (asterisk) and uniform in
texture (Figure 2F). In addition to mitochondria, cytoplasts
were also densely packed with lattice-like elements that are a
characteristic feature of mouse oocytes (Figure 2J). In many
cytoplasts examined by electron microscopy, a graded distribution of mitochondria was observed. In these instances, mitochondria appeared to be more concentrated at one end of the
oval cytoplasts. Comparisons with light microscopic images
of cytoplasts prior to thin sectioning indicated that this region
corresponded to the highly granular portion of these structures.
Human
Preliminary studies of uninseminated MII stage human oocytes
(n 5 29) subjected to a similar protocol of CCD-exposure and
centrifugation demonstrated the segregation of mitochondria
into unique compartments (Figure 3O–P). Figure 3O shows a
scanning laser confocal microscopic section midway through
an elongated oocyte after 1.5 h of centrifugation, at which
time virtually all detectable mitochondrial fluorescence
(asterisk) was localized to one portion of the cell. After 2.5 h
of centrifugation, an intense region of mitochondrial fluorescence was observed in one portion of the cytoplast (asterisk,
Figure 3P). After 12 h of culture in normal medium, cytoplast
mitochondrial fluorescence was more diffuse than observed
previously. However, a region in which mitochondrial fluorescence occurred at a comparatively high intensity could still be
detected (asterisk, Figure 3Q).
Distribution and persistence of fluorescently-tagged mitochondria introduced into intact mouse oocytes
Mitochondria were deposited in intact MII mouse oocytes
(n 5 419) by one of the following patterns of injection: (i)
deposition as bolus in the cortical cytoplasm (asterisk, Figure
3D) or at the approximate centre of the oocyte (asterisk, Figure
3F), (ii) deposition at several different sites throughout the
oocyte (asterisks, Figure 3H), and (iii) a progressive release
of aspirated mitochondria beginning at the approximate centre
of the oocyte and continuing with a progressive increase in
volume as the pipette was withdrawn (arrow, Figure 3G). The
arrows shown in Figure 3D, E, and G indicate the approximate
depth to which the injection pipette was inserted into the
oocyte. These different types of insertion were undertaken
in order to determine whether the pattern of mitochondrial
dispersion was related to the site of deposition. Injected oocytes
were cultured in microdroplets under oil with temperature
maintained at precisely 37°C in ∆T dishes positioned on a
scanning confocal microscope stage. Recipient oocytes were
examined at the initiation of culture and at 6–8 h intervals
thereafter, for up to 80 h. Optical sections were taken at
approximately the same region of the oocyte in which fluorescence from the donated mitochondria was first detected. In
Figure 1. Differential interference contrast microscopic images of intact cytochalasin D-treated GV (A, B, C; 0, 30 and 40 min after the
resumption of meiosis) and metaphase II (MII)-stage oocytes (F) after centrifugation (black asterisk indicates granular compartment; white
asterisk indicates translucent compartment). DNA fluorescence of a DAPI-stained GV-stage oocyte, and chromosomal fluorescence in a
circular bivalent (CBV)-stage oocyte and in isolated karyoplasts are indicated by arrows in D and E, and in I, J, T and U respectively. I is
a karyoplast in which the first polar body has remained attached. The asterisk in H indicates DAPI-stained mitochondria clustered to one
portion of an intact, albeit distorted MII mouse oocyte. The MII oocyte shown in F was stained for mitochondria (M; rhodamine 123, G)
and chromosomal DNA (C; YO-PRO-1-iodide, G). M, O, Q and S are cytoplasts stained with Mitotracker Red and examined in the
rhodamine isothiocyanate channel by conventional epifluorescence microscopy. The asterisks in M and O indicate areas of highest
mitochondrial fluorescence. R and S are light and fluorescent images of the same cytoplasts (black asterisks) and karyoplasts (white
asterisks). The position of the karyoplasts which do not stain for mitochondria is indicated by white asterisks in S. Chromosomal staining in
these karyoplasts is shown in T and U. K, L, N and P are light microscopic images of MII stage oocytes before (K, L) and after (N and P)
separation into cytoplasts and karyoplasts. The arrow in K and asterisk in L denote accumulation of lipid bodies at one pole of the
developing karyoplast. The asterisk in P indicates a region of increased granularity that is associated with a higher mitochondrial density.
GV, germinal vesicle; n, nucleolus; PB1, first polar body; M, mitochondria; C, MII chromosomes; L, lipid bodies (see text for further
details). (Approximate original magnifications: A, B, C, I 3700; F, G 31200; H, K, M–S 3300; T, U 3200.)
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J.Van Blerkom, J.Sinclair and P.Davis
Figure 2. Transmission electron microscopic images of centrifuged cytochalasin D-treated germinal vesicle (GV)- (A–D), GV breakdownstage mouse oocytes (G) and separated karyoplasts (E, F, H) and cytoplasts (I, J) are shown in these figures. The insert in A is a high
magnification view of the region enclosed by brackets. GV, germinal vesicle; M, mitochondria; L, lipid bodies; G, Golgi body; SER,
smooth-surfaced endoplasmic reticulum; LT, cytoplasmic lattices. (Original magnification: A, 31000 (insert, 312 000); B, 3800; C, 31000;
D, 36000; E, 31700; F, 40 000; G, 317 000; H, 314 000; I, 326 000; J, 340 000.)
some images, the original site of deposition could be recognized
as a ‘V-like’ indentation in the bolus of deposited mitochondria
(Figure 3I). This configuration appears to conform to the tip
of the pipette and was still detectable for up to 36 h in some
oocytes (Figure 3J). Mitochondria migrated from the site of
insertion in a progressive manner with the first detectable
evidence of significant movement from the bolus observed at
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~14 h (Figure 3J). For mitochondria deposited as a single
bolus near the centre of the oocyte (e.g. Figure 3F), a sphere
of fluorescence in the pericortical region of the oocyte was
observed by 36 h (Figure 3K), although a proportion of
mitochondria was still present at the original site of deposition
(Figure 3L). Cortical deposited mitochondria diffused from
the site of insertion (Figure 3M) but remained in a relatively
Interoocyte mitochondrial transfer
Figure 3. Differential interference contrast and DAPI-fluorescence images of cytoplasts (black asterisks) and karyoplasts (white asterisks)
generated by centrifugation of cytochalasin D-treated MII-stage mouse oocytes are shown in A and B. C–Q are 2 µm scanning laser
confocal microscopic images of mitochondrial fluorescence in cytoplasts derived from MII-stage CCD-treated mouse (C) and human oocytes
(O–Q). C is a pseudo-colour image. The cytoplasts shown in O, and P–Q, were stained with rhodamine 123 and Mitotracker Green
respectively. D–N are scanning laser confocal images of recipient MII stage mouse oocytes in which donated mitochondria (asterisks) were
stained with Mitofluor Green (D, H), rhodamine 123 (E, F) or Mitotracker Green (I–N). PB1 5 first polar body (see text for details).
(Approximate original magnifications: A, B, C, I 3700; F, G 31200; H, K, M–S 3300; T, U 3200.)
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J.Van Blerkom, J.Sinclair and P.Davis
confined area, as did mitochondria inserted at different cytoplasmic locations (compare Figure 3H and N). In all intact
recipient oocytes, mitochondrial fluorescence was detected
at the end of the 80 h culture period. Typically, ~20% of
injected MII oocytes lysed or degenerated after insertion
(n 5 91/419), with most of these events occurring during the
initial phase of this study as the techniques for transfer and
pipette design were being developed. In contrast, most injected
GV (n 5 28/30) and GVB/CBV stage oocytes (n 5 54/57)
lysed after manipulation and none of those which remained
intact matured to MII. Less than 15% of MII oocytes that
were intact after mitochondrial insertion fragmented during a
subsequent 80 h of culture (n 5 42/328).
ATP content of cytoplasts, untreated and recipient mouse
oocytes
With the probes used in this study, the persistence of detectable
mitochondria-specific fluorescence indicated that the inserted
organelles remained intact and viable during the 80 h culture
period. Whether the transferred mitochondria remained functional and contributed to ATP production could not be determined in living oocytes as complete cell lysis is required to
quantify ATP content. However, to minimize variability that
may exist between animals, the ATP content of unmanipulated
and injected oocytes within the same cohorts was measured.
On average, the ATP content of donor oocytes (n 5 289) was
~740 fmol (range 720–790 fmol). This value is consistent with
previous measurements of ATP contents in mature mouse
oocytes (Van Blerkom et al., 1995a). Within the individual
cohorts, injected MII oocytes (n 5 413) showed an ATP
content that was between 40 and 90 fmol higher than average,
with values .120 fmol measured in approximately one-third
of recipient oocytes (n 5 144/413). Whether these values
reflected an actual contribution by the donated mitochondria
to ATP production was difficult to determine, because within
each cohort, differences in ATP content of 30–50 fmol were
measured in the normal appearing unmanipulated MII oocytes.
However, ATP contents of over 100 recipient oocytes were
higher (800–910 fmol) than those measured in the untreated
oocytes from the same cohort(s) (690–790 fmol), suggesting
that the donated mitochondria may have increased ATP generation in these recipient oocytes.
The ATP content of mitochondria-enriched cytoplasts ranged
from 530–680 fmol (average 608 fmol, n 5 86), while the
ATP content of karyoplasts ranged from 25–51 fmol (average
39 fmol, n 5 55). The ATP content of the same volume of
cytoplasm that was injected into oocytes was determined for
78 aspirates taken from regions of cytoplasts with different
amounts of granularity/density detectable at the light microscopic level. As noted above, the occurrence of a higher
intensity of mitochondria-specific fluorescence in the more
granular regions of the cytoplasts indicated that this region
may contain mitochondria at a relatively higher density than
the less granular portions of these structures. Typically, the
average ATP content of 10 pl of cytoplasm aspirated from
cytoplasts was approximately 35 fmol. However, higher ATP
values (45–90 fmol) were derived when aspiration occurred
from the region of the cytoplasts where mitochondria appeared
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more densely packed. In contrast, ATP values ,20 fmol were
measured when aspirates were taken from regions of the
cytoplast that were presumed to have comparatively fewer
mitochondria. Similar measurements from karyoplasts were
difficult to perform because of their small size and their
tendency to lyse during manipulation. The ATP content of
nine 10 pl karyoplast aspirates was consistently ,2 fmol.
Discussion
Our earlier studies indicated that mature human oocytes can
have very different ATP contents, that the level of ATP
generation in a specific oocyte persists into the early
embryonic stages, and that oocyte-specific metabolic activity
may be associated with the developmental competence of
the embryo (Van Blerkom et al., 1995a). This study also
demonstrated that experimentally reducing ATP production in
mouse oocytes cultured in the presence of uncouplers of
oxidative phosphorylation, while not affecting fertilization
rates, did have a pronounced adverse influence on the rate at
which embryos developed to the blastocyst stage. In the human,
it has been suggested that deletions in mtDNA may occur
at high frequency in the oocytes of women of advanced
reproductive age (Keefe et al., 1995), and if the metabolic
activity of these organelles is reduced significantly or they
are unable to replicate, the effects on the embryo may be
developmental abnormality or arrest (Van Blerkom et al.,
1995a). In addition to potential metabolic consequences, mitochondrial abnormalties may also exert an influence on the
normality of pre-ovulatory oocyte maturation. In the mouse, a
mitochondrially associated trait appears to be responsible for
the inheritance of the meiosis I error Dipl I which results in
the ovulation of diploid oocytes arrested at or during the first
meiotic division (Beermann et al., 1988). This finding indicates
that mitochondrial disorders can affect the orderly segregation
of chromosomes during meiotic maturation. However, as the
central elements involved in the generation of ATP in oocytes
and embryos, any alteration in normal mitochondrial function
involving the entire complement of these organelles, or absent/
reduced activity in a significant fraction of the complement,
could have profound effects on the ability of the embryo to
develop progressively and normally.
Mitochondrial function and dysfunction in mammalian
oocytes and early embryos
Mammalian oocyte mitochondria are unique structures whose
state of development is consistent with the relatively low level
of metabolic activity associated with the ‘quiescent,’ fully
grown oocyte during meiotic arrest. In most species studied
to date, mitochondria in these oocytes are small spherical
structures containing few, undeveloped cristae that surround
an electron-dense matrix. In the mouse (Hillman and Tasca,
1969) and rabbit (Van Blerkom et al., 1973), mitochondrial
differentiation begins around the four- to eight-cell stage and
appears to involve the entire complement. Mitochondrial
differentiation is detected biochemically by increased rates
of glucose utilization, oxygen consumption, carbon dioxide
production, and net cell-specific ATP production (Mills and
Interoocyte mitochondrial transfer
Brinster, 1967; Ginsberg and Hillman, 1973; Biggers and
Borland, 1976). At the subcellular level, differentiating mitochondria are characterized by their elongation and an increased
number of cristae which penetrate a mitochondrial matrix of
reduced electron density. All of these fine structural features
are consistent with increased metabolic activity and levels of
ATP production (for review, see Van Blerkom, 1989).
While a low net ATP content appears to affect developmental competence, a similar situation may also exist when
abnormally high rates of ATP synthesis occur in early preimplantation stage embryos. For example, excessively high
rates of ATP synthesis have been detected in two- and fourcell pre-implantation lethal tw32/tw32 and t12/t12 mouse embryos,
perhaps as a result of the ability of these mutant embryos to
utilize pyruvate and lactate more efficiently than their wild
type counterparts (Ginsberg and Hillman, 1975). During the
morula stage, however, ATP levels drop below wild type
values and at least for the tw32/tw32 embryo, this drop is
associated with the development of structurally abnormal
mitochondria characterized by the presence of mitochondrial
‘crystals’ composed of divalent cations and inorganic phosphate
(Hillman and Hillman, 1975). The formation of mitochondrial
crystals is a pathological process that prevents oxidative
phosphorylation of ADP, and their occurrence in these mutant
embryos is correlated with a reduced capacity to generate ATP.
Ginsberg and Hillman (1975) suggested that a metabolic
imbalance caused by excessive carbohydrate metabolism and
ATP production in very early embryos could be responsible
for the deposition of mitochondrial crystals and the subsequent
drop in mitochondrial function as demonstrated by significantly
reduced levels of ATP generation at the eight-cell stage. This
sudden drop in ATP levels at the morula stage, following
abnormally high levels in the early cleavage stage embryo, is
a major factor contributing to the lethal phenotype expressed
in these mutants at the morula stage.
In the human pre-implantation stage embryo, mitochondrial
differentiation associated with higher levels of metabolic
activity seems to occur in a gradual fashion with increasing
numbers of mitochondria undergoing the transition from the
relatively inactive oocyte form to the more ‘orthodox’ active
form as the embryo progresses through cleavage to the blastocyst stage (Dvorak and Tesarik, 1985; Van Blerkom, 1989).
In both mouse and human, an ATP deficit may occur such that
normal stage-specific increases in biosynthetic activities (e.g.
transcription and translation) and cellular morphodynamic
processes involved in blastocyst formation, expansion, or
escape from the zona pellucida cannot be supported by the
mitochondrial complement. Indirect support for this notion is
derived from fine structural observations of human embryos
that arrested development during the pre-implantation stages
(Van Blerkom, 1989). In these cleavage-to-early blastocyst
stage embryos, mitochondrial differentiation either did not
occur or involved only a very small portion of the complement
in each cell. An association between mitochondrial activity
and normal development is also suggested by the pronounced
redistributions these organelles undergo in the maturing
oocyte and early embryo. In the mouse, for example, mitochondria are uniformly distributed in the GV stage oocyte
but are translocated (Van Blerkom and Runner, 1984) by a
microtubule-mediated process (Van Blerkom, 1991) to the
perinuclear region during formation of the first metaphase
spindle. Van Blerkom and Runner (1984) suggested that this
activity may have evolved to concentrate the limited energyproducing capacity of the oocyte to specific cytoplasmic
regions that require a higher level of ATP than would otherwise be available to support such developmental processes as
spindle assembly/disassembly, chromosomal segregation at
metaphases I and II, and polar body formation. Disruption or
inhibition of mitochondrial translocation in the mouse oocyte
was associated with perturbations in chromosomal organization
and segregation (Van Blerkom, 1991). A somewhat similar
processes of mitochondrial translocation may occur in the
newly fertilized human egg, where these organelles concentrate
around the juxtaposed pronuclei (Van Blerkom, 1989). Mitochondrial translocation to a perinuclear location also appears
to be a fundamental morphodynamic process in the early
hamster embryo (Barnett et al., 1996) and its occurrence has
been suggested to be required for developmental competence
(Barnett et al., 1997). Sousa et al. (1997) reported that
mitochondria-associated calcium may be mobilized in pronuclear human eggs and cleavage stage embryos and when
these organelles are in a perinuclear location, mobilized calcium
may be involved in signalling between the nucleus and
cytoplasm, perhaps in the regulation of the cell cycle. Consequently, the effort expended by oocytes and early embryos
to regulate mitochondrial density would seem not to be
coincidental, but rather one that reflects a normal developmental
process to adjust or provide sufficient levels of ATP or calcium
where needed within the cytoplasm. However, if the level of
mitochondrial metabolism is reduced as a consequence of
intrinsic defects (e.g. genetic) or extrinsic conditions (e.g.
oxidative stress), redistributions, translocations, or aggregations
of these organelles may not be sufficient to support a normal
process of spindle assembly or chromosomal segregation. In
this respect, Gaulden (1992) has proposed that persistent
intrafollicular hypoxia may be associated with reduced levels
of oocyte ATP production which in turn could contribute to the
generation of chromosomal segregation disorders, especially in
older women (.40 years) where such errors occur at high
frequency (Battagalia et al., 1996). A very similar situation was
reported for younger women undergoing ovarian stimulation for
in-vitro fertilization (IVF), where aneuploid oocytes with low
ATP contents were found to originate primarily from severely
hypoxic follicles (Van Blerkom, 1996; Van Blerkom et al.,
1997). Although untested, it is an intriguing possibility that a
mitochondria-associated metabolic defect subtends the chronic
production of incompetent or aneuploid oocytes observed in
some infertile women who undergo repeated failed IVF
attempts (Zenzes et al., 1992).
A direct relationship between mitochondrial function and
developmental capacity may explain an apparent, but as yet
unproven capacity to promote the developmental potential of
seemingly incompetent oocytes by the introduction of ooplasm
aspirated from the oocytes of young women (Cohen et al.,
1997). Although the precise cellular structures and macromolecules introduced into the oocyte are unknown, mito2865
J.Van Blerkom, J.Sinclair and P.Davis
chondria may represent an important component (St John and
Barratt, 1997).
Stage-specific changes in oocyte sensitivity to centrifugation
result in the generation of mitochondria-enriched cytoplasts
Cran (1987) reported that mitochondria, lipid bodies, and
membrane-bound vesicles in CCD-treated immature sheep,
pig and cow oocytes segregated into distinct zones after
centrifugation. Both the stage-related effects of centrifugation on the shape and internal organization of CCD-treated
oocytes and the separation of MII oocytes into cytoplasts and
karyoplasts described here were unanticipated. At the GV
and GVBD stages, while some differential segregation of
mitochondria and other cytoplasmic components was detected
at the fine structural level, compartmentalization of the ooplasm
into distinct mitochondria-rich and mitochondria-poor regions
was not observed, and separation of the oocyte into unique
karyoplasts and cytoplasts did not occur with the standard
conditions used in this study. However, at MII, segregation of
the cytoplasm into a chromosome-containing compartment that
was relatively devoid of mitochondria, and a chromosomefree compartment densely packed with mitochondria occurred
relatively rapidly during centrifugation and was followed by
the separation of the oocyte into two compartments, a karyoplast and a cytoplast. The mitochondria-containing cytoplast
and chromosome-containing karyoplast could be distinguished
at the light microscopic level on the basis of size, shape
and granularity. The subcellular basis for this stage-specific
response of CCD-treated oocytes to centrifugation is unknown
but may reflect normal changes in the organization and
distribution of cytoplasmic and subplasmalemmal actin filaments that occur during meiotic maturation in the mouse (Maro
et al., 1986; Van Blerkom and Bell, 1986).
Dispersion and putative activity of donated mitochondria in
recipient mouse oocytes
The findings demonstrate that the mitochondria-rich cytoplasts
retain sufficient integrity when placed in standard culture
medium to withstand micromanipulation, puncture with a
micropipette, and removal of a portion of their contents. This
result provided the basis upon which mitochondrial transfer to
intact oocytes could be contemplated.
While GV and GVBD stage oocytes remained intact after
mitochondrial insertion, virtually all underwent lysis during
subsequent culture. The transferred mitochondria remained as
a bolus at the site of insertion up to time of lysis. In contrast,
transfer-related lysis in MII oocytes occurred at low frequency,
and sequentially timed analysis of the location of fluorescently
tagged mitochondria in living oocytes by scanning confocal
microscopy showed a progressive migration of these organelles
away from the site of insertion, especially when deposition
occurred in the approximate centre of the cell. The cause of
an apparent sensitivity to microinjection and cytoplasmic
transfer in immature mouse oocytes is unclear, but it seems
unlikely that technical artefacts are responsible because oocytes
at different stages of maturity were cultured in the same
droplet and injected sequentially with the same pipette. Perhaps
stage-related differences in cytoplasmic structure and organiza2866
tion make oocytes at earlier stages of maturation less able to
tolerate injection or accommodate the transferred cytoplasm
than their counterparts of more advanced maturity. However,
it remains to be determined whether modifications to the
transfer procedure may be required in order to improve the
rate of survival of immature oocytes.
Although mitochondria remained fluorescent for the 80 h
duration of culture, suggesting that they were still viable, the
extent to which they may actually have contributed to ATP
production in the recipients could not be determined in the
living state. However, preliminary ATP content analysis of
injected oocytes during the first 36 h after cytoplasmic insertion
indicates that the donated mitochondria may increase the net
ATP content by between 40 and 90 fmol over the average
ATP content of untreated oocytes within the same cohort, and,
in ~30% of recipients, ATP content .100 fmol over the
average value were measured. Some of the ATP values
determined for recipient oocytes were higher than any ATP
content measured in an untreated oocyte from the same cohort.
At present, we consider increased ATP contents in recipient
oocytes to be suggestive rather than definitive evidence for a
metabolic contribution by the transferred mitochondria. Perhaps the multiple insertions of mitochondria may provide
unambiguous support for their contribution to oocyte metabolism. The maximum number of mitochondria that can be
transferred without compromising the integrity or developmental potential of the oocyte needs to be determined. The latter
issue is of particular relevance because of the developmentally
lethal effects of hypermetabolism observed in some early
cleavage stage mutant mouse embryos.
It is also important to emphasize that other cytoplasmic
components and macromolecules which co-migrate with mitochondria during centrifugation are also being transferred, and
their effect on development, if any, is unknown. With these
caveats taken into consideration, our findings are encouraging
with respect to the feasibility of mitochondrial donation
between oocytes, as they strongly suggest that once inserted,
the donor mitochondria remain viable and may contribute to
the ATP generating capacity of the oocyte. Studies to assess
the relative number of mitochondria transferred in a known
volume of cytoplasm, the maximum volume of cytoplasm and
number of mitochondria that can be transferred to an oocyte,
and the developmental viability of the resulting embryos are
in progress.
Potential applications of mitochondrial donation and the
issue of heteroplasmy
If reduced developmental competence of human oocytes and
embryos has a metabolic basis associated with adverse genetic
or epigenetic influences on mitochondria, then mitochondrial
donation from presumably competent oocytes as described
here may be beneficial for those gametes whose intrinsic
metabolic capacity is at a threshold between viability and nonviability. The addition of normal mitochondria in cases where
mitochondrial genetic defects are known to exist (Clayton,
1992) may be clinically relevant in the treatment of certain
maternally inherited disorders. In this instance, it may be more
appropriate to target mitochondrial transfer to a blastomere(s)
Interoocyte mitochondrial transfer
that is a progenitor of the inner cell mass, assuming that such
a cell could be identified in an early embryo (Antczak and
Van Blerkom, 1997; Gardner, 1997). At present, potential
applications of mitochondrial transfer and metabolic engineering are speculative and, to become testable, will require that
injected oocytes fertilize and develop normally, that the donated
mitochondria enter the inner cell mass lineage, and that donor
mitochondria with specific genetic markers can be identified
in the embryonic stages, as well as in fetal and adult tissues.
Heteroplasmy, the occurrence of more than one mitochondrial genotype in an organism, is rare in healthy
individuals but is associated with numerous clinical abnormalities resulting from point mutations and mtDNA rearrangements (Lightowlers et al., 1997). Heteroplasmy exists in the
newly fertilized human oocyte because the entire sperm tail is
incorporated into the ooplasm (Ash et al., 1995; Van Blerkom
et al., 1995b). However, there is no convincing evidence to
suggest that paternal mitochondria replicate or persist,
although it has been suggested that they may simply be diluted
beyond recognition during development (Ankel-Simons and
Cummins, 1996). Attempts to create a heteroplasmic state by
mitochondrial insertion into embryos have not been successful.
For example, mitochondria isolated from somatic cells and
injected into mouse early embryos were undetectable by
molecular genetic analysis in adult tissues (Ebert et al., 1989).
However, both sperm and somatic cell mitochondria are fully
differentiated, metabolically active, and, at least in the case of
the somatic cells used in these studies, were able to replicate
previously. However, the apparent inability of such mitochondria to persist does not necessarily demonstrate that
an unavoidable barrier to heteroplasmy exists. Alternatively,
insertion of fully functional, terminally differentiated mitochondria into the cytoplasm of an oocyte or embryo may be
inconsistent with the maintenance of normal metabolic activity
and replication for several days, and under such circumstances,
this condition may have an adverse influence on the ability of
the transferred mitochondria to remain functional (Corbisier
and Remacle, 1993). For the human, the approach to transfer
described here would have the advantage of both donated
and ‘recipient’ mitochondria being at the same state of
differentiation and level of metabolic activity. Whether this
factor is a critical determinant in the survival, replication, and
persistence of donated mitochondria is under investigation.
However, before any potential clinical applications could be
contemplated, the creation of a constitutive heteroplasmic
condition would have to be demonstrated in animal models,
and for therapeutic purposes the normality of the donor mtDNA
would have to be determined.
Acknowledgements
We thank Dr Samuel Alexander for his clinical contributions. This
work was supported by a grant from the National Institutes of
Health, National Institute of Child Health and Human Development
(HD31907).
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Received on April 2, 1998; accepted on July 8, 1998
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