Cells determine cell density using a small protein bound to a unique tissue-specific phospholipid

Cells and cell culture

PAT cells were isolated from 16 day chick embryos by a modification of the procedures described by Dehm & Prockop (1971). Specifically, a higher serum level (3%) was used in the dissociation medium and the use of a nylon mesh (20 µm; TETKO) instead of filter paper to separate out single cells. In these experiments the medium was changed from F12 to a 50/50 mix of F12/DMEM in order to be consistent with the medium used to grow the U2OS cells. The medium contained 0.2% fetal bovine serum deactivated for 30 m at 56°C.

U2OS cells (from frozen stocks that have been in the laboratory for over a dozen years (Campeau et al., 2009)) were grown in F12/DMEM medium with 0.2% fetal bovine serum. In 75 cm² flasks the medium, 50–75 ml, was changed every other day. When the cells first overgrow the monolayer, they are very sensitive to medium volume and will apoptose if too much medium is added.

CM was produced by growing cells in 225 cm² flasks. The medium was changed to serum-free medium and the volume reduced to 12 ml. The flasks were placed in an environmental rotator at 100 rpm, 37°C for 1 h. The CM was collected and new medium added, and the process was repeated for the third hour. After the 3rd collection medium with serum was added (50–75 ml) and the flask returned to the regular incubator.

The 16 day chick embryos were exempt from requiring an animal use protocol as determined by LBL Animal Welfare and Research Committee. Similarly, the legs from 5 week mice (BALB/c) came from mice already euthanized for other approved protocols. This is also an exempt use.

Alkaline phophatase assay

The Sigma kit was used (85L2-1KT) and the included procedures were followed with only slight modifications for labeling tissue culture flasks instead of slides.

Genes, cloning, and expression

RNA was extracted from confluent cultures of PAT cells using Trizol (Invitrogen). The RNA was reversed transcribed using superscript II (Invitrogen). The cDNA was amplified by PCR using proof reading taq polymerase (Invitrogen) using primers from the 5′ end of exon 2 plus a methionine start site and the 3′ end of exon 7.

• Forward primer: GCTGGATCCATGAAAGACCGTCTCAACCTTCCA

• Reverse primer: ACAGAATTCGACTTCATCCTGGGCTCC

This was cloned into the vector pcDNA3.1/Myc-His(+)A,B,C (Invitrogen) and sequenced. The sequence agreed with the predicted expression of this gene (minus exon 1; accession: XP_427094). Using primers for standard T7 promoter/priming site and the BGH Reverse priming site with the addition of a Sal1 site, the gene with tags was amplified using PCR. It was TOPO cloned (Invtrogen) to enhance Sal1 activity and then cut with BamH1 and Sal1. This was ligated to pESY-Neo-II (Koh, Chen & Daley, 2002; Lee, 2007), a retroviral vector, that had also been cut with BamH1 and Sal1.

In summary, at the time the sequence for the chicken exon 1 was not known, so a 3 AA sequence containing methonine was added to the 5′ end (above the dashed line below) and myc and his tags added to the 3′ end (below the dashed line below). This was inserted into a retroviral vector pESY-Neo-II – retroviral pEYK 3.1 was modified to remove the GFP and bleomycin DNA and replace it with Neo/Kan antibiotic resistance gene and a multi-cloning site (Lee, 2007) and renamed pESY1c:

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R L N L P S V L V L N S C G I T C A G D E N E I A A F C A H V S E L D L S D N K L E D W
H E V S K I V S N V P H L E F L N L S S N P L S L S V L E R R C A G S F A G V R K L V
L N N S K A S W E T V H T I L Q E L P D L E E L F L C L N D Y E T V S C S P V C C Q S L
K L L H I T D N N L Q D W T E I R K L G I M F P S L D T L I L A N N N L T T I E E S E D
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L L Q S Y T T E E R R K L L I A R L P S I I K L N G S I V G D G E R E D S E R F F I R Y
Y M E F P E E E V P F R Y H E L I T K Y G K L E P L A V V D L R P Q S S V K V E V H F
Q D K V E E M S I R L D Q T V A E L K K H L K T V V Q L S T S N M L L F Y L D Q E A P
F G P E E M K Y S S R A L H S Y G I R D G D K I Y V E P R M K
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S N S A D I Q H S G G R S S L E G P R F E Q K L I S E E D L N M H T G H H H H H H

Alternating normal and bold/italic fonts indicate exons and the underlined AAs in last exon indicate the secreted form of the gene (94 AA).

The vector pESY1c (4 µg/60 mm² dish) was used to transfect phoenix cells using lipofectamine 2000 using the manufacturer’s protocol (Invitrogen). The cells were incubated overnight and the next day medium was collected, filtered (0.45 µm), and used to transfect pESY1c into U2OS cells in the presence of 8 µg/ml polybrene. The transfected cells were selected using G418 (100 µg/ml).

Polyclonal antibody

The secreted form of pEsy1c (underlined AA above, 94 AA) with a methionine at the N-terminus (above dashed line) was inserted into the E. coli expression vector pTrcHis2-TOPO (Invitrogen). The vector included myc and his tags at the C-terminus (below lower dashed line).

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E P L A V V D L R P Q S S V K V E V H F Q D K V E E M S I R L D Q T V A E L K K H L K
T V V Q L S T S N M L L F Y L D Q E A P F G P E E M K Y S S R A L H S Y G I R D G D K
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D D D D K K G E F E A Y V E Q K L I S E E D L N S A V D H H H H H H

• Forward primer: GTGACCATGGAGCCCTTGGCAGTCGTGGAT

• Reverse primer: CTTATCGTCATCGTCCTTCATCCTGGGCTC

The procaryotic expression vector yielded an expressed protein that was 7 AA smaller in the linker between the protein and the tags (the variation is a function of the type of multiple cloning site and the primers used in cloning) and 1 AA larger at the N-terminus. This difference was too small to detect on gels and the linker region was not important for antibody production. The protein was expressed in E. coli and purified and nickel columns (Invitrogen). This protein (4 mg) was used to immunize rabbits and for affinity purification (Biosynthesis). This protein was also used as a standard for the secreted form of the factor with tags.

Immunofluorescence

Cell culture: cells were fixed with 4% paraformaldehyde for 10 m in a high calcium buffer (.05 M TES pH 7.2, 0.1 M NaCl, .05 M CaCl₂). The fixation was quenched with two, five minute washes in buffer A (.05 M TES pH 7.2, 0.1 M NaCl, .01 M CaCl₂) containing 0.1 M glycine. A blocking agent (1% casein Hammerstein [EM Science] in PBS) was used for 30 m followed by a rinse with buffer A. The primary antibody was a polyclonal made in rabbits against the secreted form of pESY1c plus tags expressed in E. coli. This was added at 1:40 (the antibody was diluted by 25% with stabilizer [Candor]) for 2 h with agitation. Then, washed 3X with buffer A with agitation and then a fluorescently tagged goat anti-rabbit secondary was added for 1 h with agitation. Washed 3X with buffer A and then once with a DAPI buffer (500 nM in PBS). Finally, a wash with buffer A.

As a control, a preimmune serum from the same rabbits was used at 1:40 (the serum was diluted 25% with stabilizer) and used as described above.

Tissue sections: hind legs from 5 week old mice were dissected to reveal the muscle and the attached tendon. These were embedded in OCT (Sakura) for 10 to 30 m and then frozen in small embedding molds in a dry ice/ethanol bath. Frozen tissue was sliced into 16 µm sections and attached to superfrost plus slides (VWR). The sections on the slide were outlined with a pap pen and then analyzed as described above. In this case, because the tendon had less autofluoresence in the red, this was the preferred color for the secondary emission. Because the plane of the knife and the plane of the tendon were infrequently in agreement, the analysis required many legs and sections.

Western blot analysis

For cytoplasmic proteins: cells were lysed (400 µl/75 cm²) with a detergent buffer (150 mM NaCl, 1% NP-40, and 50 mM Tris [pH 8.0]). This was diluted 2X sample buffer (Invitrogen) and treated as described below.

For extracellular protein: samples were concentrated on 10 kD spin ultrafilters (Millipore) diluted with 2X sample buffer and concentrated again to <200 µl. The samples were made 5% β-mercaptoethanol and heated at 80°C for 10 m. 40 µl or less was applied and run on an SDS gel (4–20%, 1.5 mm, 10 well, Tris-Glycine [Invitrogen]). The gels were blotted onto nitrocellulose (0.45 µm), washed with water, and dried overnight. Next, the blots were rehydrated with water and then blocked with 1% Hammerstein casein in PBS for 1/2 h. After 2X water washes the primary antibody (monoclonal myc (Millipore) or polyclonal rabbit, see above) was added at 1:1000 in PBS with 0.2% casein Hammerstein for 2 h. The blot was then washed 3X for 5 m in PBS with .02% Tween 20. After which the rabbit secondary antibody conjugated to horseradish peroxidase (Thermo) was added at 1:1000 in PBS with 0.2% casein. Again the blot was washed 3X for 5 m in PBS with .02% Tween 20 and then developed with a Dura or Femto kits (Thermo). The blots were photographed with a FluoroChem HD2 (ProteinSimple).

Antibody selection and specificity

Antibody affinity to a protein could be strongly affected by a lipid cofactor. This became an issue when the monoclonal antibody (clone 4A6, Millipore) to the myc tag stopped working. Over the course of these experiments Millipore changed the storage buffer several times and finally to a liquid stored at 4°C. This latter change reduced the affinity of the antibody so that only the free protein factor was detected (data not shown). The benefit of the antibody from the 4A6 clone is that it “recognizes and is specific for recombinant proteins containing the Myc epitope tag in a variety of sequence contexts”. Other clones are known to show strong affinity differences depending on the protein attached to the myc tag. Add a lipid cofactor to a protein with a myc tag and it is unclear how a context sensitive antibody would bind. To find an adequate replacement, three alternatives were tested: a rabbit polyclonal antibody to the secreted form of the factor (94 AA + tags) expressed in E. coli; a monoclonal antibody to the myc tag from Applied Biological Materials (clone A7) and one from GenScript (clone 2G8D5). Using the same CM sample for each antibody yields a Western blot that is unique to the probing antibody (Fig. 1). Critical to our purification procedure for the cofactor, only the polyclonal antibody can detect the free factor at our test concentration of ∼0.5 ng as well as bands with the factor/cofactor. Most importantly, one cannot compare the intensity of one band to that of another because each antibody has its preferred conformations; instead, one can only compare the relative amount of a specific band after different treatments.

Cofactor purification

The CM was filtered through a 0.2 µm filter and then the NaCl level was increased by adding 1/10 the volume with 5 M NaCl. The high salt CM, 10 to 50 ml, was concentrated to 1 ml in a stirred cell (10 ml; Millipore), at 700 rpm and pressurized (14 psi, N₂). The filter was a YM-10 with a nominal cutoff at 10 kD (Millipore). The retentate was diluted to 10 ml with water and reconcentrated to 0.5 ml. The retentate was analyzed on Western blots to see the level of free factor released.

The filtrate containing the cofactor was concentrated using a Pierce C18 spin column (Thermo). The column was wetted with 1 ml of 50% acetonitrile/water and then spun for 1m at moderate speed in a clinical centrifuge. Then, 1 ml of the filtrate was added and spun and repeated until all the filtrate was bound to the column. After which the column was washed twice with 1 ml of water, and twice with 1 ml of acetonitrile. Then, the washing was repeated: 1 ml of 50% acetonitrile, 1 ml water, 2X 750 µl acetonitrile, and eluted with 2X 750 µl of chloroform. Samples were stored in chloroform and concentrated before analysis by blowing down with nitrogen to ∼100 µl. For mass spectrometry, 10 µl of sample was diluted with 80 µl of acetonitrile and 10 µl water. All solvents were lc, ms, or hplc grade.

With recently purchased YM-10 filters (in comparison to using older filters from the original maker Amicon), the cofactor would not separate from the factor unless the following changes were made: the filters had to soak at 37°C in 20% acetonitrile for 48 h; after concentrating the high salt CM to 1 ml, it was put into 10 ml of buffer (10 mM EDTA[pH 8.0], 2 ml acetonitrile, 3 ml isopropyl alcohol, 5 ml water) and heated to 80°C for 2 h and then concentrated to 0.5 ml using the treated YM-10 filter. The filtrate was further concentrated and purified as described above.

Lipid enzyme treatment

Lipid degradative enzymes were first used to characterize the makeup of the cofactor. In this case, we were looking for a change in the mobility of factor/cofactor on a Western blot after being treated with these enzymes for various times 30 m, 90 m, 180 m, and a no enzyme control. CM (3 ml) was concentrated to 0.5 ml (one sample for each time point). Then the enzymes were added as detailed below and incubated for the specific time, 2X sample buffer was added to stop the reaction, concentrated on a 10 kD spin ultrafilter, and analyzed by Western blotting.

• Lipase: Sigma (L8525); 380 units/µl; added 50 µl; incubated 37°C in buffer B (see below)

• Phospholipase C: Sigma (P7633); .038 units/µl; added 50 µl; incubated 37°C in buffer B

• Phospholipase A₂: Sigma (P9279); 6 units/µl; added 50 µl; 20 mM triethanolamine pH 8.9 substituted for TES in buffer B; incubated room temperature.

• Sphingomyelinase: Sigma (S9396); 0.54 units/µl; added 2 µl; incubated 37°C in buffer B with MgCl₂ substituted for CaCl₂.

The second use of lipid enzymes was to identify the peaks on mass spectrometry that were sensitive to the enzyme treatment. In the cofactor purification above, between the wash steps, the column material was resuspended in buffer B (20 mM TES pH 7.2; 10 mM NaCl; 1 mM CaCl₂), enzyme added, and incubated for 30 min at 37°C with agitation. This was spun down and the protocol above continued. In the buffer, when using sphingomyelinase, CaCl₂ was replaced with MgCl₂.

Mass spectrometry analysis

The diluted lipid extract samples were analyzed on an ABSciex TripleTOF 5600 (AB Sciex, Foster City, CA). Samples were introduced to the mass spectrometer via syringe pump by using a Nanospray III source (ABSciex) with a nano-tip emitter (New Objective, Woburn, MA) operating in positive-ion mode (2400 V). The data were acquired with Analyst TF 1.5.1 by averaging the signal from 400 m/z to 1250 m/z over several minutes. MS/MS spectra were collected in “high sensitivity” mode with the quadrupole set to UNIT resolution and collision energy set to 30 to optimize fragmentation. MS/MS spectra were scanned from 100 m/z to 1600 m/z and were collected for a total accumulation time of 500 ms.

Expression in U2OS cells causes differentiation

A replication defective retroviral vector containing a neomycin resistance gene was used to transfect the chicken gene plus tags into U2OS cells (Koh, Chen & Daley, 2002; Lee, 2007). Because this study was started when the chicken genome was first sequenced, the chicken equivalent to the first exon was not present. So the insert contained only the last six exons (pESY1c).

U2OS cells, derived from a human osteosarcoma tumor, showed stable expression of this gene after infection when grown in low serum media (0.2%) (Valmassoi & Schwarz, 1988). Despite the low serum levels, U2OS cells transfected with pESY1c would start to overgrow the monolayer as small ridges. The ridges would coalesce into circles sharing boundaries with other circles that resembled spongy bone in a two dimensional space (Fig. 2B). Uninfected U2OS cells showed some ability to overgrow the monolayer but could not maintain the growth needed to form completed structures (Fig. 2A). The multicell ridges produced alkaline phosphophatase on the cell surface, indicating increased expression of the osteoblast phenotype (Fig. 2C).

Pattern of molecular expression intracellularly and extracellularly

A monoclonal myc antibody and Western blotting was used to probe the expression and processing of this myc tagged gene. Two specific bands were seen intracellularly: ∼44 kD band which is expected for the full size gene product; ∼26 kD band that could be a cleavage product (Fig. 3). However, this band is larger that the predicted 20 kD protein (16 kD + tags).

To observe the secreted forms a simple purification was used whereby the CM was put through a 30 kD ultrafilter. Both the retentate and the filtrate were analyzed and both had to be further concentrated on a 10 kD ultrafilter in order to load sufficient protein on the gel for Western blot analysis (Fig. 3). The retentate contained many bands: the smallest was a weak band at ∼28 kD, the strongest was at ∼60 kD, and the largest was over 200 kD. The filtrate showed basically a single band at ∼60 kD. This was unusual since 60 kD proteins are excluded to a high degree from passing through a 30 kD ultrafilter. One explanation is that by concentrating the sample ∼20-fold causes the tagged protein to aggregate into stable forms that resist the denaturing effects of heat, β-mercaptoethanol, and SDS.

Aggregation is due to a cofactor

The multiple bands seen in the Western blot complicates the interpretation of how this tagged protein is working in the extracellular space. But more specifically, even the smallest MW band (∼28 kD) seen outside the cell is higher than predicted. Is this larger size due to a different amount or type of modification in bone cells versus tendon cells? Or is this increase in apparent MW a reflection of an interaction with another molecule?

To begin to answer these questions, we needed to resolve how the unmodified secreted form of the protein factor with two tags (94 AA + tags) migrates on a Western blot. To accomplish this, the tagged protein was expressed in E. coli, purified on a nickel column, and then used as a standard in Western blots. The tagged secreted form of the protein ran with a MW of 16 kD (Fig. 4, lane 6). This confirms that there is either a covalent modification or a binding to another molecule that gives rise to the 28 kD form in U2OS cells. Further aggregation could then occur when concentrating the factor to give rise to larger MW forms.

This raises the question of whether different physical or chemical environments could alter the size of the bands observed on Western blots. CM from U2OS-pESY1c was put through a 30 kD ultrafilter and the filtrate was then concentrated on a 10 kD ultrafilter. This was divided into 2 parts. One was further concentrated on a 10 kD ultrafilter for loading on a gel but in this case, 30% DMSO was added. To the other sample, salt was added to raise the concentration by 0.5 M; concentrated with an ultrafilter; then diluted to low salt; and then concentrated to apply to a gel (Fig. 4). The effect of DMSO was to reduce the number of bands, especially those at high MW, but the smallest band still remained 28 kD. In contrast, the treatment with high salt, low salt and ultrafiltration yielded a much simpler picture where the bands were reduced to the 16 kD band and its dimer at 32 kD. One can conclude that the 28 kD band is due to a binding of a cofactor and not a covalent modification. In addition, when the cofactor is separated from the secreted form of the protein factor, the MW of the protein is the expected 16 kD (94 AA + tags). This shows that U2OS cells are capable of cleaving the large gene product to the 94 AA + tags and then secrete it.

Localization and characterization of the factor/cofactor in the extracellular space

The previous experiment raises the question of how the untagged 16 kD form in tendon cells compares to the 28 kD tagged form seen in bone cells. Even taking into account a 4 kD tag, there is an additional 8 kD shift in MW. Are these forms variations with common characteristics or is the bone form acting in a different manner? In tendon cells the 16 kD form is loosely bound to the cell layer. This gives a key feature to this form in that gently shaking a flask of tendon cells causes them to loose their ability to sense high cell density, resulting in increased proliferation.

First, the localization was examined with immunofluorescence using a rabbit polyclonal antibody specific to the tagged secreted form of the chicken protein expressed in E. coli. During tests to optimize the signal, procedures that disrupted membranes decreased the signal. Using an optimized protocol, immunofluorescence staining was localized to areas where the cells have overgrown the monolayer (hills) and expression was suppressed in monolayer areas (valleys) (Fig. 5A).

Second, the cells were subjected to gentle shaking (100 rpm) for 3 h per day with the medium changed each hour. After 3 days the hills had spread into the valleys and the valleys had additional growth. The net effect was to even out the growth over the flask (Fig. 5C).

One can conclude that some of the major physical properties of the bone and tendon factor/cofactor are the same. They are both localized to the cell layer – most likely bound to the cell membrane. However, this interaction is not strong, allowing significant diffusion that can be enhanced by gentle agitation of the medium.

Characterization of the bone cofactor with degradative enzymes

The cofactor is tightly bound to the factor and gives a consistent mobility pattern on Western blots. This pattern could then be used to test whether specific degradative enzymes could cleave the cofactor, alter the migration profile, and reveal the partial composition of the cofactor.

One possible candidate for the cofactor would be a variation on the anchoring structure for membrane bound proteins: a glycosylphosphatidylinositol lipid (Paulick & Bertozzi, 2008). To test for a glycolipid, the factor/cofactor was treated with a variety of polysaccharide degradative enzymes – chondroitinase ABC, hyaluronidase, heparinase – and the pattern on a Western blot was compared to that of an untreated control. None of the treated samples showed a distinct change in the pattern (data not shown). At the same time, a variety of lipid degrading enzymes were tested – lipase, phospholipase A₂, phospholipase C, and sphingomyelinase. Lipase, phospholipase C, and sphingomyelinase caused dramatic shifts in the pattern seen on Western blots (Fig. 6). Lipase was the most dramatic causing bands to shift to lower MW including the 16 kD, free protein form. The lipase was a crude preparation used at a high concentration; so while its primary target would be a glycerol fatty acid linkage in the cofactor, this is probably not the only cleavage product. The phospholipase C used in this experiment has overlapping activities with the sphingomyelinase. The sphingomyelinase is more specific to cleaving moities on ceramide phosphates. In contrast, phospholipase A₂ did not show a significant change in mobility or intensity (data not shown). A negative result is hard to interpret because there are too many potential causes. The positive results, while not yielding a definitive structure, implicate lipid components containing a phosphoceramide and a glycerol fatty acid as part of the cofactor. The lipid nature of the cofactor can explain why the factor/cofactor has a strong tendency to aggregate when concentrated due to hydrophobic interactions.

Purification and further characterization of the bone cofactor using mass spectrometry

In order to characterize the bone cofactor in more detail, procedures were needed to separate both the factor from the cofactor and then the cofactor from other lipid-like molecules. The first step in the purification – the separation of the cofactor from the factor – could benefit from standard lipid procedures using a methanol/chloroform extraction. In this procedure the lipids are found in the chloroform layer and proteins are found as a precipitate between the layers. There is no assay for the cofactor but the proteins in the precipitate can be solubilized and then Western blot analysis can be used to see if the protein factor runs as the free form (16 kD) or as a complex with the cofactor (28 kD). The amount of free factor is an indicator of cofactor release. However, redissolving the protein precipitates at the interface and running Western blots only showed the 28 kD form and higher order aggregates (Fig. 7). In this extraction protocol, the cofactor remained bound to the factor, and as a consequence, precipitated at the interface.

Despite attempts to weaken the interaction between the factor and the cofactor by changes in pH, salt, solvent, chaotropic agents, temperature, and chelating agents, Western blots showed little or no conversion to the free form (data not shown). The only successful method required a combination of a shift in salt concentration and ultrafiltration (Fig. 4). Using a simplified version of this procedure whereby the CM was made high in salt (∼0.6 M) and then concentrated on a 10 kD ultrafilter. This was then diluted 10-fold with water and concentrated to run on Western blots (Fig. 8). By intensity of the bands, most of the tagged factor runs as the free form. Thus, the corresponding free cofactor was expected to have passed through the ultrafilter as part of the filtrate.

Using a small C18 resin column to concentrate, differential solvent elution to purify, and then comparing the untreated and the enzyme treated samples via mass spectrometry one can identify candidate cofactor peaks (Fig. 9). Only molecular weights between 530 and 580 are shown because there was no significant differences between the samples at either higher or lower values (mass range = m/z 300 – m/z 1500). Two doubly-charged ions at m/z 543.4273 and m/z 557.4429 were dramatically reduced by treatment with either enzyme and these peaks were not observed in the buffer only control and the intensity of the m/z 557 ion was highly correlated with the intensity of the free protein factor in the retentate. The observed difference between the doubly-charged ions was 14 representing a 28 Da difference in MW commonly seen for lipids. In the rest of the analysis, we focused on the 557 species with a MW of 1114.8858. By using a high mass accuracy Q-TOF mass spectrometer we generated potential molecular formulas with a list of atoms commonly found in organic biomolecules (i.e., carbon, hydrogen, nitrogen, oxygen, and phosphorous). The most likely formula C63H123N2O11P (exact mass = 557.4432252+; 0.6 ppm from the measured mass) had nearly a 1:2 Carbon:Hydrogen ratio common to lipids and the nitrogen, oxygen and phosphorous indicative of a hydrophilic region. Support for the proposed cofactor structure derived from MS/MS fragmentation of m/z 557. Two fragment ions, shown in Fig. 10, at m/z of 323 and m/z 378 were consistent with a ceramide phosphate and a linker moiety, supporting the results from the enzymatic reactions.

From our analysis, the cofactor was susceptible to three lipid enzymes – lipase, phospholipase C, and sphingomylenase, and a peak observed at m/z 557 that has a strong lipid composition. This peak was lost when treated with lipase, phopholipase C, and sphingomylenase. Using our knowledge of which enzymes cleave the cofactor and the molecular formula, a working model can be postulated that includes a ceramide phosphate, a glycerol lipid, and a linker consisting of a short modification of a proline residue between them (Fig. 10).

Tendon cofactor characterization using enzymes and mass spectrometry

Having analyzed the bone factor/cofactor, the focus returned to the original aim of characterizing the tendon factor, but now knowing that it binds a cofactor. The bone and tendon cofactors must be different because they impart a unique shift in mobility to the factor/cofactor complex on SDS gels. This raises important questions about the nature of the tendon cofactor and how it compares to its bone counterpart.

As a first step in the characterization, the same panel of lipid degradative enzymes were tested on PAT cell CM. In this case, concentrated CM was used that was over 6 months old. This results in a stable aggregate at ∼60 kD being the dominant band. As with bone, lipase and phospholipase C were active on the PAT cell factor/cofactor while phospholipase A₂ was inactive (Fig. 11A). Unlike the bone cofactor, sphingomyelinase was inactive on the PAT cell factor/cofactor (Fig. 11B). To insure that the inactivity of the enzyme was not due to the factor/cofactor being in a specific aggregate, the treatment with sphingomylinase was repeated with fresh CM (Fig. 11C) and the result remained unchanged. This indicates a lack of a ceramide phosphate moiety in the tendon cofactor.

The cofactor was purified for mass spectrometry by binding it to C18 columns followed by washing with methanol, acetonitrile, and elution with chloroform. Via mass spectrometry we compared preparations that were treated with either phosphospholipase C or lipase while the cofactor was bound to the C18 column to the untreated cofactor samples. No differences were seen in the methanol or acetonitrile washes; however, the chloroform fraction did reveal several peaks that changed between the untreated and treated samples. Two singly charged peaks, at 668.61 and 696.65, were reduced to background levels by the enzyme treatments. Similar to the bone cofactor mass spectrometric analysis, these peaks differed by 28 MW and this reflects a variation in lipid chain length by two carbons. Additionally, there was no doubly charged ion observed at m/z 557 as was seen in the bone cofactor sample (data not shown).

Using MS/MS to further characterize the 668.61 ion yielded a rather simple spectrum with two peaks of roughly equal intensity and with MW of 283.26 and 311.29. These ions differ by 28 and accurate mass measurements correspond to saturated fatty acids with 18 and 20 carbons, respectively. Both chains are required to be in the model in order to achieve the high lipid content indicated by the parent molecule’s MW. Since the band is degraded by phospholipase C, there is a phosphate group yet the total mass of the intact ion is insufficient to include a commonly observed glycerol phosphate. Consequently, our proposed structure includes anhydride linkage of the fatty acid directly to the phosphate. The mass difference between the total mass and the fatty acid and phosphate groups corresponds to a cyano group, thus our working model adds a cyano group to the phosphate and is consistent with the enzymology, the mass spectrometry data, and the gel mobility data (Fig. 12). Whether the cyano group was part of the original molecule or it displaced another group during the purification remains to be determined.

Nomenclature

With two different cofactors and the potential for more in the future, there is a need to give them distinct names. For the bone factor/cofactor will be named SNZR (sensor)1P1L; for tendon factor/cofactor will be named SNZR 1P2L. Similarly, the protein factor will be SNZR 1P and the bone cofactor, SNZR 1L and the tendon cofactor, SNZR 2L. At present, because of the extremely high homology between species at the protein level and no knowledge of the conservation of lipid structure between species, the delineation is only on cell type.

Cofactor tissue specificity

The cofactor for bone and the one for tendon are unique. The SNZR 1P can bind either SNZR 1L or SNZR 2L and drive a unique differentiation pathway dependent on the type of cofactor produced by the cell. This raises the question of whether the SNZR 1P2L when added to U2OS cells is capable of driving bone differentiation or does it elicit a new or null response? In addition, one can ask whether the protein factor, by itself, has any activity?

To answer these questions, U2OS cells were grown in regular media as a control, or with 5X concentrated condition media from U2OS-pESY1c cells expressing the SNZR 1P1L, and from 5X concentrated condition media from PAT cells expressing SNZR 1P2L. In addition, SNZR 1P purified from E. coli, was added at 5 µg/ml – 3 orders of magnitude greater than the estimated level in CM (Figs. 13A–13D). These levels were chosen because early work with PAT cells have shown that 1X CM is insufficient to stimulate growth when added back to confluent cultures (Zayas & Schwarz, 1992). The high level of the protein factor was chosen because it seemed unlikely that the factor without the cofactor would have a lot of activity and small amounts of activity are difficult to detect by visualization.

After a week in the various media, the most dramatic effect was that the tendon CM was inhibitory to growth when added to the U2OS cells yielding a subconfluent culture. The addition of the SNZR 1P alone did not appear to change the cultures compared to the control even at this high concentration. Adding 5X CM from U2OS cells expressing the SNZR 1P1L, showed some increased in growth as the cells appeared to be more compact. However, to confirm this observation we allowed this flask and its control to grow for 9 additional days, at which point it was obvious that the CM had caused the cells to completely overgrow the monolayer (Figs. 13E, 13F). What is particularly striking is the uniformity of growth when the CM is added to cells versus the lattice patterns that are formed when the cells themselves express the SNZR 1P1L (Fig. 2B).

Tissue localization in mice

One aim of this paper – producing the recombinant form of the tendon cell density signal in another cell type and then applying it to tendon cells to show it has the original activity – is not compatible with our data. Since the cofactor that is essential for activity is unique (in two cases) to the cell type producing it, another test would be useful in showing that the protein factor is the correct candidate for being part of the cell density signaling mechanism. There is the similarity that both the tendon SNZR 1P2L and bone SNZR 1P1L are loosely bound to the cell layer and cause the cells to differentiate. In addition, the polyclonal antibody to the 94 AA protein does detect a band at 16 kD in the PAT cell CM (Fig. 11C). A more functional test would further confirm the identity.

To accomplish this, the polyclonal antibody was used to check whether the factor was at the tendon/muscle junction of 5 week old mice. This is the growing stage for mice and one would predict that the factor would be at high concentration at the growing ends of the tendon where there should also be a high density of fibroblasts (Schwarz, 1996). To detect the protein factor, a polyclonal rabbit antibody was used with a secondary antibody labeled with alexa 568; to detect the nuclei, the sections were stained with DAPI (Fig. 14).

At the growing end of the tendon there is a strong fluorescence signal for the factor (Fig. 14A) as well as numerous nuclei labeled with DAPI (Fig. 14B). There is variation in the signal over the length of the tendon but this can result from the plane of the tendon not being in exactly the same plane as the section. The strong staining for the factor at the growth plate of the tendon supports its role as a cell density signal.

As a control, a preimmune serum was used at an equal concentration and exposed for equal time and it is basically negative (Fig. 14C) and the DAPI staining shows the tendon nuclei starting at the lower left corner and rising steeply to the top of the picture (Fig. 14D).

One unexpected observation is the light staining of the muscle connective tissue with its single file nuclei (Figs. 14A and 14B). This is an indication that a third cell type may be producing this protein factor along with its own unique cofactor.