J. Embryol. exp. Morph. 86, 205-218 (1985)
Printed in Great Britain © Company of Biologists Limited 1985
205
Transferrin in foetal and adult mouse tissues:
synthesis, storage and secretion
JENNIFER MEEK AND EILEEN D. ADAMSON
La Jolla Cancer Research Foundation, 10901 North Torrey Pines Road, La
Jolla, California 92037, USA
SUMMARY
Transferrin is an important growth-promoting serum glycoprotein synthesized chiefly in the
liver in adults. The transferrin found in the mouse foetus is thought to be wholly a product of
the foetus itself and its synthesis starts at least as early as the 7th day of gestation. The major
sites of synthesis in mouse foetuses are the visceral yolk sac (VYS) and liver (Adamson, 1982).
We now report that other murine foetal tissues synthesize readily detectable amounts, namely
lung, spleen, spinal cord and rib cage. Very low levels are also synthesized by the brain, muscle
and pancreas. We can detect no synthesis of transferrin in late foetal thymus, heart or skin
although mid-gestation foetal skin may make a very small amount. No synthesis of transferrin
can be detected in adult brain, lung and spleen, but approximately equal rates of synthesis are
detected in adult liver and adult ear pinna.
Transferrin is accumulated by foetal and adult tissues in widely varying amounts and these
have been measured by enzyme-linked immunosorbent assays of extracts. In addition to VYS
and liver, high levels of transferrin are found in foetal skin, lung and rib cage with lower
amounts in spinal cord, spleen and muscle tissues. Tissues of the 15th day foetus accumulate
the highest concentrations of transferrin. A role for the mediation of transferrin in the stimulation of growth and differentiation by interacting tissues is discussed.
INTRODUCTION
Transferrin is a j3i-globulin glycoprotein (Mr = 80 000) present in serum at
2 - 4 m g m r 1 (Putnam, 1975), and capable of tightly binding two ferric and other
metal ions. Its principal function is regarded as the transport of iron, and its targets
are those tissues which are actively growing such as embryonal and tumour cells
and those cells with special iron requirements such as haematopoietic cells. Iron
(and zinc) is required for all dividing cells to maintain enzyme and co-factor levels
in new cells. Transferrin is known to be adsorbed to cells through a specific receptor
glycoprotein (Mr = 180000) and is probably endocytosed into vesicles called
receptosomes. From here an acidic compartment removes iron and this is found
later in the cytosol, bound to ferritin. Transferrin, unlike most ligands which enter
via receptors, appears to be recycled out of the cell and reutilized in the extracellular fluid (reviewed by Octave, Schneider, Trouet & Crichton, 1983).
Embryonal cells have special demands for transferrin for the synthesis of ironcontaining haem and enzymes. Since there appears to be a barrier to the passage of
Key words: Mouse, transferrin, glycoprotein, liver, yolk sac.
206
JENNIFER MEEK AND EILEEN D. ADAMSON
transferrin from the maternal circulation (Faulk & Galbraith, 1979) at least up to
the 18th day of gestation (Renfree & McLaren, 1974), it follows that transferrin
should be an early product of the murine embryo. The earliest time that transferrin
synthesis has been detected (Adamson, 1982) is on the 7th day of gestation. The
endoderm layer of the two-layered egg cylinder appears to be the source and later
this layer develops into the endoderm of the visceral yolk sac (VYS). The VYS and
the foetal liver are major sources of transferrin during gestation. Both of these
endodermally derived tissues also synthesize alphafoetoprotein (Dziadek & Adamson, 1978), but only transferrin continues to be synthesized by the liver in adults.
It is not yet known if transferrin is synthesized by any other foetal mouse organs,
an important question in view of the ability of transferrin to stimulate the proliferation and differentiation of mouse foetal kidney (Ekblom et al. 1983), chick muscle
(Markelonis, Oh, Eldefrawi & Guth, 1982a; Ii, Kimura & Ozawa, 1982; Beach,
Popiela & Festoff, 1983), and lymphocyte growth in vitro (Brock, 1981; Anderson,
Chase & Tomasi, 1982). Transferrin is an essential component of serum-free media
(Sato et al. 1979) for all kinds of cell types. We have therefore made a survey of
murine foetal tissues to identify those that synthesize transferrin and which may
stimulate their own development or the development of associated tissues.
We have analysed the content of transferrin in several adult and foetal organs in
order to identify transferrin which may be sequestered or accumulated to significantly high levels. The hypothesis is that transferrin could be released locally or
preferentially to influence the development of associated tissues. These data
together with transferrin synthesis rates are assessed to determine the possibility of
a role for transferrin in tissue interactions which are known to stimulate organogenesis or cell proliferation.
MATERIALS AND METHODS
Metabolic labelling and extraction of foetal tissues
Random bred mice (CDI Flow Laboratories) were mated and the day offindingthe copulation
plug was designated thefirstday of gestation. Pregnant females were killed by cervical dislocation,
the uteri removed and foetuses dissected to collect the tissues. Care was taken to remove skin and
brain samples before opening the body cavity and dissections were made so that the liver fragments
did not contaminate any other tissue. Tissues (0-05-0-1 g wet weight) were washed in TrypsinEDTA for 10-30 min and twice in medium (DME with 10 % foetal bovine serum) before preincubating them in 0-5 ml DME containing 1/20 normal concentration of methionine and 10 %
dialysed foetal bovine serum. After 30min, [35S]methionine (lOOjuCiml"1, l-2Ci i umor 1 , New
England Nuclear), in fresh medium was added and incubation continued at 37 °C in 7 % CO2 in air.
After 3 h, tissues were separated from the incubation medium, and the latter was centrifuged at
15000 g for 10 min at 4 °C. Tissues were washed once in phosphate-buffered saline and homogenized
in 0-5 ml hypotonic buffer. Lysing buffer contained 10mM-KCl, lmM-MgCl2, pH7-5, 0-1 mMPMSF, and after 15 min at 0 °C, Triton X-100 was added to 0-5 %. After centrifugation at 15000gfor
10 min the supernatant was removed for analysis and the pellet was dissolved in 0-5 ml 0-5 N-NaOH
to assay for protein by the Coomassie blue-dye-binding procedure (Sedmak & Grossberg, 1977).
The medium and extracts were analysed as shown in theflowdiagram (Fig. 1) for protein, total
and acid-insoluble radioactivity, total transferrin content by enzyme-linked immunosorbent
assay (ELISA) (see below) and for immunoprecipitated radioactive transferrin (see below).
Transferrin synthesis and secretion
207
Dissected tissue 3h in
medium + [ 3S S|methionine
Tissue
Medium
Aliquots
for TCA ppt.
Aliquots
for ELISA
Hypotonic buffer, 0-5 7c Triton,
inhibitors, ccntrifugation
15000/?, 20min.4°C
Most for
immunoppt
& PAGE
Dissolve in 0-5 N NaOH
Aliquots for
protein assay
Aliquots for
TCA ppt.
Aliquots
for ELISA
Most for
immunoppt
& PAGE
Aliquots for
protein assay
Fig. 1 Flow diagram to show foetal tissue extraction and analysis procedures.
Analysis of radioactive protein
From the medium and extracts prepared as above, 2jul aliquots were examined for total
radioactive protein by precipitation on filter paper squares with ice-cold 10 % (w/v) trichloracetic
acid (TCA) (30min at 0°C). The squares were washed with cold 10 % TCA containing 1 mMmethionine, followed by ethanol. They were dried and counted in 3 ml Beta-Max (West Chem
Products, San Diego, CA) in a scintillation counter.
Appropriate aliquots of medium and extract were taken to contain 2 X 105 and 4 x 105 c.p.m.
TCA-insoluble radioactivity, respectively. Aliquots of medium were made up to 1 to 1-2 ml with
NET buffer (0-4M-NaCl, 5mM-EDTA, 005M-Tris-HCl pH8, 1% NP-40, 0-lmM-PMSF and
0-02 % sodium azide) and one portion was treated with 5 [A (approx. 5 ]Ug) affinity-purified antitransferrin (see below) while to another, 5 [il non-immune rabbit immunoglobulin was added.
After incubation at room temperature for lV^h or overnight at 4°C, 30 jul of a 50 % suspension
of protein A-agarose (Sigma) was added and the mixture mixed end-over-end for lVzh at 4°C.
The beads containing adsorbed antibody/antigen complexes were washed twice in NET buffer
and once in 0-0625 M-Tris-HCl pH6-8 before being boiled in 30//1 two-fold-concentrated gel
sample buffer (Laemmli, 1970). Tissue extracts were analysed similarly except that the diluting
and washing medium was RIPA buffer (0-01 M-Tris-HCl, pH7-5, 0-15 M-NaCl, 1 % NP-40,1 %
sodium deoxycholate, 0-1 % sodium dodecyl sulphate). Radioactive proteins were analysed by
electrophoresis on 7-5% polyacrylamide gels (Laemmli, 1970), stained with 0-2% (w/v)
Coomassie brilliant blue, destained and soaked influor(Autofluor, National Diagnostics, Somerville, NJ), dried and exposed to preflashed Kodak XAR-5filmfor 5-21 days at - 70 C (Bonner
& Laskey, 1974). The bands on films were analysed by scanning densitometry and quantitated
by integration of the peaks.
Analysis of transferrin by ELISA
Pooled mouse plasma was processed by the method described by Sawatzki, Anselstetter &
Kubanek (1981) to isolate pure transferrin. Briefly, a 45% saturated-ammonium-sulphatesupernatant fraction from mouse serum was salted out on a column of Sepharose CL-6B, and the
transferrin peak was rechromatographed on DEAE-Sepharose. Peak fractions from this column
were homogeneous by several criteria (Adamson, 1982). Transferrin (200 fig) dissolved in PBS
was mixed with an equal volume of Freund's complete adjuvant and injected into several subcutaneous sites in one rabbit. Immunizations were repeated at 4-week intervals using Freund's
incomplete adjuvant. Serum from the third bleed onwards contained anti-transferrin activity in
208
JENNIFER MEEK AND EILEEN D. ADAMSON
double-diffusion tests. Antibodies were affinity purified by passing a fraction which was
precipitated by 40 % saturated ammonium sulphate over a conjugate of 50 mg purified transferrin linked to cyanogen-bromide-activated Sepharose (10 ml). After washing away unadsorbed
protein, antibodies were eluted with O-lM-glycine-HCl, pH2-6. Fractions with the highest
precipitating activity were pooled, dialysed against 0 05 M-NH4HCO3 solution and lyophilized.
The antibody was shown to react specifically with mouse transferrin by double-diffusion
analyses (Adamson, 1982) and by lack of crossreaction with whole mouse, calf and rabbit sera
in ELISA.
A portion (1 mg) of affinity-purified anti-transferrin antibodies was crosslinked to 5 mg alkaline
phosphatase (bovine intestine, Sigma type VII) according to the method described by Engvall
(1980) and this was used as the third layer in a sandwich ELISA. The wells of 96-well polystyrene
plates (Linbro/Titertek, cat. no. 76-38104, Flow Laboratories, Inc., McLean, VA) were coated
with affinity-purified rabbit anti-mouse transferrin (100 fA of 5 fig. ml"1 antibodies dissolved in
OlM-sodium bicarbonate buffer at pH9-5). This was allowed to adsorb for 16 h at room temperature. The plate was washed three times with 0-9 % (w/v) NaCl, 0-05 % (v/v) Tween 20 after
each layer. The second layer was a solution (100 jul) containing an unknown or standard transferrin concentration from 2 to 200ng.ml"1 in PBS, 5% calf serum, 0*05% Tween-20 (Bovine
transferrin does not react with the anti-mouse transferrin antibodies, Adamson, 1982, and
therefore transferrin in calf serum does not interfere in the assay). Incubation and washing
followed as described above. Rabbit antimouse transferrin coupled to alkaline phosphatase
(200 jul of 0-5/ig. ml"1) was added to each well for 5h at room temperature and finally, 250 (j\
substrate for alkaline phosphatase (Sigma 104 at 1 mg. ml"1 in 0-1 M-diethanolamine-HCl, 1 mMMgCl2, pH9-8) allowed the enzyme activity in each well to be revealed as a yellow colour. The
absorption at 405 nm was measured in a Flow titertek 96-well reader after 20 min and 30 min of
colour development. Figure 2 shows a typical standard curve used to measure the concentration
of transferrin in tissue extracts. The lower limit of sensitivity is 10 ng. ml"1 transferrin.
Measurement of transferrin in adult tissues
Adult mice were perfused by passing 0-9 % NaCl solution into the heart of anaesthetized
heparin-injected animals using a 20 cm pressure head. Tissues were dissected and approximately
1 gm was homogenized and processed as described for foetal tissues.
5
10
20
40
100 200
Transferrin concentration (ng. ml"1)
Fig. 2 Standard curve for transferrin ELISA. The concentrations of transferrin in
individual wells (averages of duplicates) of a 96-well plate are shown on the abscissa (log
scale). The optical densities of the products of alkaline phosphatase-linked (third layer)
anti-transferrin antibodies is shown on the ordinate.
Transferrin synthesis and secretion
209
RESULTS
Transferrin synthesis by foetal tissues
Figure 3 A shows that large amounts of transferrin are synthesized by the liver and
VYS of foetuses. Smaller amounts of radiolabelled transferrin are secreted into the
medium by spinal cord (including ganglia), spleen, rib cage and lung. Some transferrin is also synthesized by the brain, skeletal muscle and pancreas. In general,
transferrin synthesized at very low levels is detected best in the medium after the
3h incubation period, while the tissue extract contains much less. Transferrin
synthesis was undetectable in 19th day thymus, heart and skin. Several experiments
gave varying results and these are given separately in Table 1. Variations may arise
in part because of differences in the exact stage of development between litters and
in part by differential resistance to damage during processing. Some tissues of 15th
day foetuses were also examined. Low levels of transferrin synthesis were detected
in the spinal cord, brain and skin with large amounts in VYS incubates or 15th day
tissues (Table 1).
It is clear that several foetal tissues can synthesize and release transferrin into the
medium. However, examination of Coomassie-blue-stained gels showed that
synthesis of transferrin may be of only minor importance compared to its storage
or retention by certain tissues. Figure 4 shows that the skin of 19-day foetuses
secretes sufficiently high levels of both alphafoetoprotein (lane 1) and transferrin
(lane 2) after immunoprecipitation to stain as distinct protein bands on gels.
Table 1. Detection of transferrin synthesis in foetal tissues by metabolic labelling
15-day foetus
Tissue
VYS
liver
S.C.
brain
thymus
spleen
rib cage
muscle
heart
pancreas
lung
skin
medium
+
19-day foetus
tissue*
medium
tissue'
-
+
++
-, -, -
—
++
-, -
VYS = visceral yolk sac
S.C. = spinal cord
* Tissue and incubation media were analysed separately for immunoprecipitated transferrin.
f For extracts that gave variable results, individual experiments are displayed between commas. Blank spaces indicate the tissue was not examined. The number of + signs indicates
approximate quantitation by densi tome try.
210
JENNIFER MEEK AND EILEEN D. ADAMSON
Similarly, spinal cord (lane 4), brain (lane 6), ribs (lane 11), muscle (lane 13) and
heart (lane 15), in addition to VYS (lane 8) and liver (lane 10) release readily
detectable transferrin into the medium during the 3 h incubation period. We therefore measured the transferrin content of various foetal tissues at several stages.
Transferrin content of foetal and adult tissues
To establish the size of the compartment of retained transferrin, total transferrin
1 2 3 4 5 6 7 8
_
MrX
910
1 2 3 4 5
m
200_
-TF
92.56843-
(A)
_
_
(0
1 2
3 4
1 0 -3
5 6
M rX
10 -3
200 -
20092.5-
68-'
43-
(B)
'
1 2
3 4
Transferrin synthesis and secretion
211
in tissue extracts and incubation media was measured by ELISA. These values are
expressed as /ig transferrin. ing"1 protein in Figure 5 and as jug. g"1 wet weight of
tissue in Table 2. Among tisues of the 13th day foetuses, the placenta holds the
M
x
10 -3
M 1 2 3 4 5 6 7 8 9
1 0 1 1 1 2 1 3 14 15
^
200 -
-TF
«**
Fig. 4 Coomassie-blue-stained polyacrylamide gel showing the products analysed after
immunoprecipitation of 19th day foetal tissues. Lanes 1-10 are the same as Figure 3A;
lane 11, ribs, anti-transferrin; lane 12, skeletal muscle, control; lane 13, skeletal muscle,
anti-transferrin; lane 14, heart, control; lane 15, heart, anti-transferrin. MT markers are
shown on the left and the positions of transferrin (TF) and alphafoetoprotein (AFP) on
the right. Specifically immunoprecipitated protein bands are indicated by spots on the
left hand side.
Fig. 3 Autofluorographs of immunoprecipitated metabolically labelled extracts.
[35S]methionine was incorporated into tissues in organ culture for 3 h and media or tissues
were immunoprecipitated with anti-alphafoetoprotein, anti-transferrin or preimmune
rabbit control Ig as described in Materials and Methods. (A) Immunoprecipitated
products from the medium of 19th day foetal tissues: lane 1, skin, anti-alphafoetoprotein;
lane 2, skin, anti-transferrin; lane 3, spinal cord, control; lane 4, spinal cord, antitransferrin; lane 5, brain, control (this precipitate appears to have missed the washing
steps but serves to indicate the range of total biosynthesized proteins); lane 6, brain, antitransferrin; lane 7, VYS, anti-alphafoetoprotein; lane 8, VYS, anti-transferrin; lane 9,
liver, anti-alphafoetoprotein; lane 10, liver, anti-transferrin. (B) From 19th day foetal
tissues: lane 1, lung, anti-transferrin; Lane 2, lung, control; lane 3, spleen, antitransferrin; lane 4, spleen, control; lane 5, pancreas, anti-transferrin; lane 6, pancreas,
control. (C) Immunoprecipitated products from the medium of 18th day foetal tissues:
lane 1, spinal cord, control; lane 2, spinal cord, anti-transferrin; lane 3, VYS, antitransferrin; lane 4, liver, anti-transferrin; lane 5, ribs, anti-transferrin. (D) adult tissues:
lane 1, ears, control; lane 2, ears, anti-transferrin; lane 3, liver, control; lane 4, liver, antitransferrin. The positions of relative molecular mass (Mr) markers are shown on the left
and transferrin (TF) and alphafoetoprotein (AFP) on the right. Lower Mr bands in the
heavy immunoprecipitates obtained from VYS and liver are likely to be breakdown
products in these tissues; the higher Mr band seen in (B), lanes 5 and 6 and in (C), lane 5, is
fibronectin which is specifically adsorbed by protein A. Exposures were for 14-38 days.
212
JENNIFER MEEK AND EILEEN D. ADAMSON
20 -
15
I
I
10
i
I
2 > _J CQ
13th
15th
19th
Tissues (day of development)
Fig. 5 Transferrin content of foetal and adult tissues. Transferrin was assayed in tissue
extracts by ELISA and is expressed as /ig. mg"1 tissue protein. Conditions are given in
Table 2. Tissues are: M, muscle or limb-bud; V, visceral yolk sac; L, liver, R, ribs; B,
brain; S, skin; H, heart; Lu, lungs; S.C., spinal cord; K, kidney; P, placenta; T, thymus;
Sp, spleen; Pa, pancreas; S.N., sciatic nerve; S.G., salivary gland; Ts, testes.
greatest store of transferrin, expressed as jug. mg" 1 protein. Considering the large
size of the placenta compared to all other tissues, the importance of this store is
clear. Up to the 15th day of gestation, the VYS appears to be predominant over the
liver, but later in gestation the two tissues become equally important sources of
transferrin.
Comparisons of the transferrin content of tissues dissected from 13th day, 15th
day and 19th day foetus and adult perfused tissues show (Figure 5 and Table 2): (1)
Tissues of the 15th day foetus, namely VYS, liver, spinal cord, brain, vertebrae or
ribs, limbs and heart, accumulate higher concentrations of transferrin than any
other stage examined. In all tissues except lung, peak levels of transferrin are found
Transferrin synthesis and secretion
213
Table 2. Transferrin content of washed foetal and perfused adult mouse tissues
Tissue
15th day foetus
i"g • g" 1
wet wt.
19th day foetus
J"g • g" 1
wet wt.
775 ± 625
599 ± 506
VYS
liver
100 ± 22
197
S.C.
brain
S.N.
14 ±8
27 ±2
49 (5-173)
29 ± 9
thymus
spleen
rib cage
43-3
20
63
125 (28-278)
muscle
heart
92-3
9-2
110 ± 82
71 ± 1 6
pancreas
kidney
lung
sal. gl.
testes
11-5
28
191
128 (41-215)
skin
15-3
406 ± 232
adult
jwg • g" 1
wet wt.
352 ± 224
28
143 ± 101
170 (35-285)
114 ± 19
145 ± 36
359 ± 71
132
139 ± 47
The transferrin content of extracts was estimated by ELISA (see Methods section for details).
Washed or perfused tissues were incubated for 3 h in culture media before separate analyses of
total transferrin that was present in the medium and in the tissue. These figures were also used
to derive those in Figure 5. VYS, visceral yolk sac; S . C , spinal cord; S.N., sciatic nerve; sal. gl.,
submandibulary salivary gland.
Means of 2-4 experiments ± standard deviations with at least triplicate samples. In some cases,
parentheses showing the range are given. Single value indicates the result of one experiment with
at least duplicate samples. Blank spaces indicate no determinations.
in the 15th day foetus; (2) In the lung, transferrin continues to accumulate throughout
gestation and into the adult stage where levels become similar to the stores in liver; (3)
Nervous tissues (spinal cord and brain) of the foetus accumulate transferrin to
moderate levels which decline after the 15th day. The adult sciatic nerve contains considerable concentrations of transferrin; in some individuals this level is higher than in
the liver when expressed as a proportion of the total protein concentration in that
tissue; (4) The total store of transferrin in skeletal muscle is very high in adults and late
foetuses considering that a high proportion of body mass is muscle. In addition, 15th
day limb (predominantly developing muscle and cartilage) contains the highest
concentrations of transferrin of any tissues examined. The rib cage and vertebrae
of mid- to late-gestation mouse foetuses also contain high levels.
Release of transferrin by foetal tissues
We made a further comparison to determine the rates at which tissues could
release transferrin into the medium during organ culture. Figure 6 shows that most
214
JENNIFER MEEK AND EILEEN D. ADAMSON
15
10
I
1
C/3
Tissues (day of development)
Fig. 6 Transferrin released by dissected tissues incubated in organ culture for 3 h at
37 °C. Transferrin was determined by ELISA as described in the Materials and Methods
section. Tissues are those listed in Figure 5; data given in Figure 5 were obtained in the
same experiments.
15th day foetal tissues (muscle/limbs, ribs/vertebrae, brain, skin and heart) are
capable of releasing transferrin into the incubation medium at rates higher than that
of liver. Similar observations hold for 19th day foetal muscle, rib cage, brain, skin,
heart, lung and kidney. Very slow rates of release are detected from spinal cord,
thymus, spleen, and pancreas when expressed as fig released. mg" 1 tissue protein.
Transferrin synthesis by adult tissues
In addition to foetal liver and VYS, mouse spinal cord, brain, skin, lung, spleen,
rib cage and pancreas in foetuses from the 15th to the 19th day of gestation
synthesize transferrin as shown by metabolic labelling and electrophoretic analysis
of immunoprecipitates (Fig 3A,B,C and Table 1). If we are to conclude that these
sources of newly synthesized transferrin are significant for the development or
maturation of tissues, we might expect that they are switched off in adult tissues
when they would no longer be needed. Adult mouse lung, spleen, brain, liver and
pinna of the ear (skin and cartilage, etc.) were labelled with p5S]methionine as
described in Materials and Methods. Autofluorographs show that synthesis of
Transferrin synthesis and secretion
215
transferrin is undetectable in lung, spleen and brain, but is occurring at
approximately equal rates (expressed as c.p.m. g"1 wet weight) both in liver and in
ear pinna (Fig. 3D). The latter is a surprising and rich source that could derive from
the sebaceous glands, epidermis, connective tissue, cartilage or blood elements
contained within the pinna. Further studies will be needed to answer this question.
DISCUSSION
Synthesis of transferrin
We report here that transferrin may be synthesized by a variety of cell types in
the foetal mouse, since several tissues secrete a metabolically labelled polypeptide
that is antigenically related to adult mouse transferrin. It has previously been
reported that several adult mammalian tissues (liver, leucocytes, macrophages,
Sertoli cells, salivary gland, testis, ovary and lactating mammary gland; Thorbecke
et al 1973; Imrie & Mueller, 1968; Tormey, Imrie & Mueller, 1972; Haurani,
Meyer & O'Brien, 1973; Skinner & Griswold, 1980 and 1983) synthesize transferrin
but no foetal tissues other than liver and visceral endoderm have been identified to
have this ability in human (Gitlin & Perricelli, 1970), rat (Yeoh & Morgan, 1974)
and mouse foetuses (Adamson, 1982). Recently, however, it has been reported that
chick embryo spinal cord synthesizes transferrin (Stamatos, Squicciarini & Fine,
1983). We find that midgestation (15th day) foetal spinal cord, brain and skin
synthesize low but detectable levels of transferrin while very large amounts are
produced by the VYS and liver (Table 1). Late foetal spinal cord continues to
synthesize transferrin as do several other tissues (Fig. 3). The list of foetal tissues
that synthesize transferrin may be much larger since it is possible that our detection
system is insufficiently sensitive or that tissues are damaged during trypsin-EDTA
or other treatments. Of those adult tissues examined, only the liver and the pinna
of the ear synthesize readily detectable amounts (Fig. 3D). These results suggest
that multiple synthetic sources of transferrin are required for the special demands
of certain foetal tissues during gestation.
Importance of the visceral yolk sac
The VYS has long been known to play an important role in acting as a protective
barrier, as a transporter of selected molecules as well as producing nutritive
materials to the developing foetus. Transferrin is one of several nutritive
macromolecules synthesized very early in mouse development (7th day) by the
visceral endoderm and later by the VYS (Adamson, 1982). Other VYS products
with carrier and nutritive roles are AFP (Yeoh & Morgan, 1974; Dziadek & Adamson, 1978) and apolipoprotein (Shi & Heath, 1984). The major source of transferrin
for the rapidly proliferating cells of the developing embryo is clearly the VYS, since
the foetal liver may only commence significant synthesis by the 13th day or later
(15th day in rat, Yeoh & Morgan, 1974). The immense importance of the VYS in
supporting embryonic development cannot be overstated. It therefore may seem
216
JENNIFER MEEK AND EILEEN D. ADAMSON
that other tissues that synthesize transferrin must be relatively minor sources.
However, the proximity of a transferrin source or delivery by a special mechanism
may be important for some developing tissues especially since arteries and blood
capillaries would not yet be organized in the primordial organ buds of the embryo.
Retention of transferrin by tissues
In perfused tissues, most of the serum transferrin has been removed and what
remains is present in these compartments: (1) Intracellular transferrin newly
synthesized in that tissue; (2) Transferrin that has been endocytosed or that is
bound to cell surface receptors; (3) Transferrin in the interstitial fluid and remaining
serum. All of the compartments may release transferrin readily to the surrounding
medium. The time taken for intracellular transferrin to be released after synthesis
is less than H h (our unpublished observations) and it has been documented that
endocytosed transferrin may be released intact within minutes of entry into cells
(Octave, Schneider, Trouet & Crichton, 1983). In these studies we have measured
the release of transferrin from these combined compartments in order to estimate
the influence that each tissue may have on nearby target tissues in terms of providing an essential nutrient or stimulator, namely, transferrin.
Transferrin is found in foetuses at levels far in excess of those in adults, therefore we
conclude that transferrin is as necessary to embryonic development in general as it is
to cells cultured in defined media. Transferrin is present in 15-day foetal tissues, at a
time when rapid organ growth is occurring, at two- to fifty-fold higher levels than in
13th or 19th day foetuses. These findings indicate (a) possible specialized roles in
certain tissues such as muscle, bone and skin, and (b) the importance of a store of
transferrin at stages when the circulatory system is not well developed.
A developmental role for transferrin
Transferrin is a required factor for the proper development of muscle (Cohen &
Fischbach, 1977; Podlewski et al. 1978; Kuromi, Gonoi & Hasegawa, 1981; Markelonis etal. 19S2a,b; Stamatos etal. 1983; Matsuda, Spector & Strohman, 1984a;
Matsuda, Spector, Micou-Eastwood & Strohman, 19846), kidney (Ekblom, Thesleff, Miettinen & Saxen, 1981; Ekblom et al 1983), teeth (Partanen, Thesleff &
Ekblom, 1984) and blood (Broxmeyer et al. 1980; Pelus, Broxmeyer, de Sousa &
Moore, 1981; Gentile & Broxmeyer, 1983). We show here that transferrin levels
are highest when the foetal tissues' proliferative demands are very great and conclude that developmental requirements could be met by one or all of the following:
(a) synthesis in the individual tissue; (b) retention and reuse; (c) delivery from
adjacent tissues; (d) delivery by nerves.
There is some evidence that transferrin exerts a mitogenic effect independently
of its ability to supply iron to proliferating tissues (Imrie & Mueller, 1968; Tormey
et al. 1972). The recent findings that there is some homology between the oncogene
Blym and transferrin (Goubin et al. 1983) also support the notion that transferrin
may be a general mitogen.
Transferrin synthesis and secretion
217
We are indebted to Drs. A. Grover, S. Edwards, and D. Mercola for constructive criticisms of the work. We thank G. Sandford for excellent photography and D. Lowe for typing
the manuscript. This study was supported by grants from the NIH (HD18782, P30 CA 30199 and
CA 28427).
REFERENCES
E. D. (1982). The location and synthesis of transferrin in mouse embryos and
teratocarcinoma cells. Devi Biol. 91, 227-236.
ANDERSON, W. L., CHASE, C. G. & TOMASI, T. B. (1982). Transferrin support of stimulated
lymphocytes. In Vitro 18, 766-774.
BEACH, R. L., POPIELA, H. & FESTOFF, B. W. (1983). The identification of neurotrophic factor
as transferrin. FEBS Lett. 156,151-156.
BONNER, W. M. & LASKEY, R. A. (1974). Afilmdetection method for tritium-labelled proteins
and nucleic acids in polyacrylamide gels. Eur. J. Biochem. 46, 83-88.
BROCK, J. H. (1981). The effect of iron and transferrin on the response of serum-free cultures of
mouse lymphocytes to Concanavalin A and lipopolysaccharide. Immunology 43, 387-398.
ADAMSON,
BROXMEYER, H. E., DE SOUSA, M., SMITHYMAN, A., RALPH, P., HAMILTON, J., KURLAND, J. I. &
BOGNACKI, J. (1980). Specificity and modulation of the action of lactoferrin, a negative feed-
back regulator of myelopoiesis. Blood 55, 324-333.
A. & FISCHBACH, G. D. (1977). Clusters of acetylcholine receptors located at identified
nerve-muscle synapses in vitro. Devi Biol. 59, 24-38.
DZIADEK, M. & ADAMSON, E. D. (1978). Localization and synthesis of alphafetoprotein in postimplantation mouse embryos. /. Embryol. exp. Morph. 43, 289-313.
EKBLOM, P., THESLEFF, I., MIETTINEN, A. & SAXEN, L. (1981). Organogenesis in a defined
medium supplemented with transferrin. Cell Differ. 10, 281-288.
EKBLOM, P., THESLEFF, I., SAXEN, L., MIETTINEN, A. & TIMPL, R. (1983). Transferrin as a fetal
growth factor: Acquisition of responsiveness related to embryonic induction. Proc. natn. Acad.
Sci., U.S.A. 80, 2651-2655.
ENGVALL, E. (1980). Enzyme immunoassay ELISA and EMIT. Meth. Enzymol. 70, 419-439.
FAULK, W. P. & GALBRAITH, G. M. P. (1979). Trophoblast transferrin and transferrin receptors
in the host-parasite relationship of human pregnancy. Proc. Roy. Soc. London B. 204, 83-97.
GENTILE, P. & BROXMEYER, H. E. (1983). Suppression of mouse myelopoiesis by administration
of human lactoferrin in vivo and the comparative action of human transferrin. Blood 61,
982-993.
GITLIN, D. & PERRICELLI, A. (1970). Synthesis of serum albumin, prealbumin, a-foetoprotein,
a-antitrypsin and transferrin by the human yolk sac. Nature 228, 995-997.
GOUBIN, G., GOLDMAN, D. S., LUCE, J., NEIMAN, P. E. & COOPER, G. M. (1983). Molecular
cloning and nucleotide sequence of a transforming gene detected by transfection of chicken Bcell lymphoma DNA. Nature 302, 114-119.
HAURANI, F. E., MEYER, A. & O'BRIEN, R. (1973). Production of transferrin by the macrophage.
/. Retic. Soc. 14, 309-316.
Ii, I., KIMURA, J. & OZAWA, E. (1982). A myotrophic protein from chick embryo extract: its
purification, identity to transferrin and indispensability for avian myogenesis. Devi Biol. 94,
366-377.
IMRIE, R. C. & MUELLER, G. C. (1968). Release of a lymphocyte growth promoter in leukocyte
cultures. Nature 219, 1277-1279.
KUROMI, H., GONOI, T. & HASEGAWA, S. (1981). Neurotrophic substance develops tetrodotoxinsensitive action potential and increases curare-sensitivity of acetylcholine responses in cultured
rat myotubes. Devi Brain Res. 1, 369-379.
LAEMMLI, U. (1970). Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680-685.
MARKELONIS, G. J., OH, T. H., ELDEFRAWI, M. E. & GUTH, L. (1982a). Sciatin: a myotrophic
protein increases the number of acetylcholine receptors and receptor clusters in cultured
skeletal muscle. Devi Biol. 89, 353-361.
COHEN, S.
218
JENNIFER MEEK AND EILEEN D . ADAMSON
G. J., BRADSHAW, R. A., OH, T. H., JOHNSON, J. L. & BATES, O. J. (19826). Sciatin
is a transferrin-like polypeptide. /. Neurochem. 39, 315-320.
MATSUDA, R., SPECTOR, D. & STROHMAN, R. C. (1984a). There is selective accumulation of a
growth factor in chicken skeletal muscle. I. Transferrin accumulation in adult anterior
latissimus dorsi. Devi Biol. 103, 267-275.
MATSUDA, R., SPECTOR, D., MICOU-EASTWOOD, J. & STROHMAN, R. C. (19846). II. Transferrin
accumulation in dystrophic fast muscle. Devi Biol. 103, 276-284.
OCTAVE, J.-N., SCHNEIDER, Y.-J., TROUET, A. & CRICHTON, R. R. (1983). Iron uptake and
utilization by mammalian cells. I. Cellular uptake of transferrin and iron. Trends in Biochemical Science 8, 217-220.
PARTANEN, A. M., THESLEFF, I. & EKBLOM, P. (1984). Transferrin is required for early tooth
morphogenesis. Differentiation 27, 59-66.
MARKELONIS,
PELUS, L. M., BROXMEYER, H. E., DE SOUSA, M. & MOORE, M. A. S. (1981). Heterogeneity
among resident murine peritoneal macrophages: Separation and functional characterization of
monocytoid cells producing granulocyte-macrophage colony stimulating factor (GM-CSF) and
responding to regulation by lactoferrin. /. Immunol. 126, 1016-1021.
PODLEWSKI, T. R., AXELROD, D., RAVDIN, P., GREENBERG, L., JOHNSON, M. M. &SALPETER, M.
M. (1978). Nerve extract induces increase and redistribution of acetylcholine receptors on
cloned muscle cells. Proc. natn. Acad. ScL, U.S.A. 75, 2035-2039.
PUTNAM, F. W. (1975). In The Serum Proteins (ed. F. W. Putnam), Vol. 1, pp. 265-316. New
York: Academic Press.
RENFREE, M. B. & MCLAREN, A. (1974). Foetal origin of transferrin in mouse amniotic fluid.
Nature 252, 150-161.
SATO, G. H. et al. (1979). In Methods in Enzymology (ed. W. B. Jakoby & I. H. Pastan), Vol.
58, p. 89. New York: Academic Press.
SAWATZKI, G., ANSELSTETTER, V. & KUBANEK, B. (1981). Isolation of mouse transferrin using
salting-out chromatography on Sepharose CL-6B. Biochim. biophys. Acta 667, 132-138.
SEDMAK, J. J. & GROSSBERG, S. E. (1977). A rapid, sensitive and versatile assay for protein using
Coomassie brilliant blue G250. Analyt. Biochem. 79, 544-552.
SHI, W.-K. & HEATH, J. K. (1984). Apolipoprotein expression by murine visceral yolk sac endoderm. /. Embryol. exp. Morph. 81, 143-152.
SKINNER, M. K. & GRISWOLD, M. D. (1980). Sertoli cells synthesize and secrete a transferrin-like
protein. /. biol. Chem. 255, 9523-9525.
SKINNER, M. K. & GRISWOLD, M. D. (1983). Multiplication stimulating activity (MSA) can
substitute for insulin to stimulate the secretion of testicular transferrin by cultured Sertoli cells.
Cell Biol. int. Rep. 7, 441-446.
STAMATOS, C , SQUICCIARINI, J. & FINE, R. E. (1983). Chick embryo spinal cord neurons
synthesize a transferrin-like myotrophic protein. FEBS Lett. 153, 387-390.
THORBECKE, G. J., LIEM, H. H., KNIGHT, S., COX, K. & MULLER-EBERHARD, U. (1973). Sites of
formation of the serum proteins transferrin and hemopexin. J. din. Invest. 52, 725-731.
TORMEY, D. C , IMRIE, R. C. & MUELLER, G. C. (1972). Identification of transferrin as a lymphocyte growth promoter in human serum. Expl Cell Res. 74,163-169 and 220-226.
YEOH, G. C. T. & MORGAN, E. H. (1974). Albumin and transferrin synthesis during development
in the rat. Biochem. J. 144, 215-224.
{Accepted 15 November 1984)