The following reagents were used in this study: PCL (average molecular weight Mn = 45.00 kDa; Sigma-Aldrich, CA, United States), a-DMEM, DMEM/F12 (HyClone, UT, United States), fetal bovine serum (FBS; Gibco, NY, United States), TRITC and FITC phalloidin (R415 and A12379, Invitrogen, CA, United States), 4,6-diamidino-2-phenylindole (DAPI, C0060, MaoKang, Shanghai, China), cell counting kit-8 (CCK-8; Dojindo, Kumamoto, Japan), primary antibodies anti-collagen type II (COL II; ab185430), anti-collagen type I (COL I; ab6308), anti-aggrecan (AGC; ab3778), SOX-9 (SOX9; ab76997; Abcam, Cambridge, United Kingdom), Alexa Fluor 555-conjugated anti-mouse antibody (A32727, Invitrogen, CA, United States), RNeasy mini kit (Qiagen, Hilden, Germany), SuperScript^TM III Reverse Transcriptase (Bimake, Shanghai, China), and Sirius red solution, Safranin, and fix-green solution (YifanBio, Shanghai, China).

In this study, PCL was used as the scaffold material that was placed in the charging barrel. The MEW 3D printer (TL-Direct Writing 150) was started, and the barrel temperature was adjusted to 65°C. The distance between the needle and the printing table was about 3 mm, and the printing station had a negative electrical charge. Furthermore, the printing voltage changed as the printing height changed (6 kV, voltage; 1,500 mm/min, printing speed; 22°C, ambient temperature; 35%, humidity). Additionally, the print needle model was 21G, with 100 kPa of air pressure. The conditions remained stable during printing. Different pore supports were fabricated according to the drawing model, and each scaffold was 200 μm in thickness (10 layers).

Three-week-old New Zealand white rabbits were euthanized to extract bone marrow mesenchymal stem cells (BMSCs), chondrocytes (CCs), and tendon stem cells (TCs). For BMSC isolation, both ends of the femur and tibia were removed. Subsequently, 10 ml of high glucose α-DMEM medium containing 10% FBS, 1% penicillin/streptomycin, and 100 U/ml heparin sodium was used to repeatedly rinse the bone marrow cavity and elute the bone marrow. Furthermore, stem cells were collected in a centrifuge tube and centrifuged at 100 × g for 5 min. The supernatant was discarded, and the pellet was resuspended in high-glucose α-DMEM medium with 10% FBS and 100 U/ml penicillin/streptomycin. Finally, the cells were incubated in an incubator at 37°C and with 5% CO₂, and the medium was changed every 2–3 days.

For CC isolation, the femur and tibia were removed, and a scalpel and scissors were used to separate the articular cartilage and cut the cartilage into pieces (<0.1 mm). These pieces were suspended in 10 ml DMEM/F12 medium containing 1% collagenase II (2275MG100, Biofroxx, Germany) and mixed. The suspension was then placed in a shaker with 37°C constant temperature and digested at 100 r/min for 2 h. Furthermore, the mixture was centrifuged at 100 × g for 5 min, followed by the removal of the supernatant, and cultured in DMEM/F12 medium containing 10% FBS, 50 μg/ml ascorbic acid, and 100 U/ml of penicillin/streptomycin in an incubator at 37°C with 5% CO₂. The medium was changed every 2–3 days.

For TC isolation, the Achilles tendon was removed and cut into pieces (<0.1 mm). Subsequently, the tendon pieces were suspended in 10 ml DMEM/F12 medium containing 1% collagenase I (1904MG100, Biofroxx, Germany), blown, and mixed. The suspension was placed in a constant temperature shaker at 37°C and digested at 100 r/min for 2 h. The mixture was centrifuged at 100 × g for 5 min, and the supernatant was discarded. Finally, the cells were cultured in DMEM/F12 medium containing 10% FBS and 100 U/ml of penicillin/streptomycin in an incubator at 37°C with 5% CO₂. The medium was changed every 2–3 days.

The cells were observed daily. The extracted primary cells were marked as P0, and the P2 phase cells were used in the experiments.

After the scaffolds were printed, they were immersed in 75% alcohol and placed in an ultraviolet disinfection box to be disinfected for about 12 h. After removal, the scaffolds were rinsed with PBS three times (5 min). The scaffolds were then placed in six-well plates or a 96-well plate. Then, P2 BMSCs, CCs, and TCs were adjusted to a density of 3,000 cells/ml; 1 ml of cell suspension/well was added to six-well plates, and 100 μl of cell suspension/well was added to 96-well plates. After a 4-h incubation, the six-well plates were supplemented with 1 ml of medium per well, and the 96-well plates were supplemented with 100 μl of medium per well.

The pore size and fiber diameter of the scaffold were printed using a pre-designed model. The printed scaffold was mounted on aluminum foil and sprayed with 6-nm-thick platinum (OPC-80T, SPI Supplies, West Chester, PA, United States). The sample with cells was fixed using glutaraldehyde for 4 h and then fixed with citric acid for 2 h. After fixation, it was dehydrated using ethanol (50, 70, 85, 90, and 100%) and then placed in a lyophilizer (Free Zone, Labconco, United States) overnight. The dried sample was coated with platinum. After coating with a 6-nm-thick layer of platinum, scanning electron microscopy (Leica, Germany) was used to observe the scaffold.

Scaffolds with cells were fixed using 4% paraformaldehyde. After 1 h, the specimens were removed, washed with PBS three times for 5 min, immersed in Triton X-100 (0.1%) for 5 min, and washed three times with PBS (5 min each). Furthermore, the specimens were incubated with FITC-labeled phalloidin for 45 min at 21°C and then washed with PBS three times (5 min each). Next, the specimens were incubated with DAPI for 10 min at room temperature and then washed with PBS three times (5 min each). Finally, fluorescence microscopy (Leica, Germany) was used to observe the cells after staining.

The CCK-8 test was used to measure the adhesion and proliferation of different cells in various pore scaffolds. After the scaffolds were placed in a 96-well plate, BMSCs, CCs, and TCs were seeded on each of the pore scaffolds. At 4, 7, 14, and 21 days, the cell scaffolds were transferred to another 96-well plate. Then, 10% CCK-8 reagent was added to each well and placed in an incubator at 37°C for 1 h in the dark. Subsequently, 100 μl of each well medium was transferred to a clean 96-well plate. The absorbance of the culture media was measured with a microplate reader (Thermo Fisher Scientific, Inc.) at 450 nm (630 nm as a reference, n = 4).

P2 BMSCs were used to study chondrogenesis, 1 ml of cell suspension (3,000 cells/ml) was added to each well of the six-well plate containing scaffolds. After the cells adhered to the scaffold, the medium was replaced with chondrogenic differentiation medium (DMEM, 1% insulin–transferrin–selenium, 100 μg/ml sodium pyruvate, 40 μg/ml proline, 10^–7 M dexamethasone, 50 μg/ml ascorbic acid, and 10 ng/ml TGF-β1). The cell condition was closely observed, and the medium was changed every 2 days.

After 21 days, the specimen were removed, washed two to three times with PBS, and fixed with 4% paraformaldehyde for 1 h. Sirius red, Safranine O, and Fast-Green staining were performed separately. For Sirius red staining, the sample was immersed in PBS and washed. Then, the specimens were stained with Sirius red for 10 min, washed with ethanol, and observed under the stereoscope. For Safranine O and Fast-Green staining, the sample was immersed in PBS and washed. Subsequently, Fast-Green was added for 15 min and washed with PBS, followed by incubation with red dye solution for 5 min and treatment with color separation solution for 20 s. After PBS washing, the specimens were observed under a stereoscope.

The scaffolds with cells were fixed with 4% paraformaldehyde. After 1 h, the specimens were removed, washed with PBS three times for 5 min, immersed in Triton-X (0.1%) for 5 min, and washed three times with PBS for 5 min each. The specimens were blocked with BSA for 1 h and primary antibody (COL II, COL I, AGC, and SOX9; diluted according to the manufacturer’s protocol) for 12 h at 4°C and then washed with PBS three times for 5 min each. Subsequently, they were incubated with the Alexa Fluor 555-conjugated anti-mouse antibody for 1 h at room temperature, washed three times with PBS (5 min each), and then incubated with phalloidin and DAPI as described above in fluorescence staining and microscopy. The specimens were observed by confocal microscopy (Leica, Germany) after staining. ImageJ (National Institutes of Health, Bethesda, MD, United States) was used for quantitative analysis of the image (n = 4).

Quantitative reverse transcriptase-polymerase chain reaction (qRT-PCR) was used to detect chondrogenic differentiation-associated gene expression. Cells were seeded and stimulated using the abovementioned methods. After induction culture for 21 days, the medium was discarded, and total RNA was extracted with RNeasy mini kit (Qiagen) according to the instruction manual. The mRNA expression of the cartilage-specific genes (AGC, COL I, COL II, and SOX9) was determined by qRT-PCR using β-actin as an internal reference; the primer sequences are shown in Supplementary Table 1. RT-PCR was performed by reverse transcribing 1 mg of RNA with SuperScript^TM III Reverse Transcriptase, followed by PCR with SYBR Premix Ex Taq II (2×) on an ABI 7500 Fast machine (Applied Biosystems, Courtaboeuf, France). Gene expression was calculated using the 2^–ΔΔCt method, where ΔΔCt = (the average value of the gene Ct to be tested – the average value of the reference gene Ct to be tested) – (the average value of the target gene Ct of the control – the average value of the reference gene Ct of the control) (n = 5).

All samples were collected and weighed, and then they were tested for total collagen and type II collagen content. Total collagen content was quantified by hydroxyproline assay, and type II collagen content was quantified by sandwich ELISA in accordance with a published method (n = 5) (Reddy and Enwemeka, 1996; He et al., 2017, 2018).

Data analysis was performed using SPSS 25.0 software (SPSS Inc., Chicago, IL, United States). Quantitative data are expressed as mean ± standard deviation (mean ± SD). Data were analyzed by independent-sample t-test and one-way analysis of variance. P < 0.05 was considered significant.

Figure 2 shows the morphology of the scaffolds under a scanning electron microscope. In this study, we prepared a MEW scaffold with high precision. The surface of the printing fiber is smooth, the diameter of the wire is about 10 μm, and the pore size of the scaffold is controllable. These are conducive to follow-up research.

Bone marrow mesenchymal stem cells, TCs, and CCs were used in a comparative study of the effect of pore size on cell adhesion and proliferation. For different cells, the most suitable pore size for its adhesion and proliferation might be different. Figure 3A shows the DAPI-labeled nuclei (blue) and the phalloidin-labeled β-actin (red) in BMSCs under a fluorescence microscope and merged images. Figure 3B shows that BMSCs exhibited adhesion and proliferation on 50-, 100-, 200-, 300-, and 400-μm-pore-size scaffolds at 4, 7, and 14 days, respectively. The cell adhesion growth state in each image is significant, indicating that the scaffold has good biocompatibility and is suitable for cell adhesion. At 4 days, the cells in the 50-μm-pore scaffold were almost filled with pores. In the other groups, cell adherence to the surface of each fiber of the scaffold was observed, and there was a tendency to fill the inner pores. At 7 days, the 50- and 100-μm scaffolds were covered and filled by cells; the 200-, 300-, and 400-μm scaffolds also showed a trend of cell ingrowth, which was significantly increased compared with 4 days. At 14 days, the five groups of scaffolds were filled. In each group, compared to 4, 7, and 14 days, the number of cells increased significantly. Among them, the number of cells filled in the 50 μm group increased, and the nuclear field of DAPI significantly increased. Figure 3C shows the CCs’ adhesion and proliferation on 50-, 100-, 200-, 300-, and 400-μm-pore-size scaffolds at 4, 7, and 14 days. Compared to BMSCs at 4 days, chondrocyte adhesion occurred faster, and the number of adherent cells was higher. At 7 days, the chondrocytes spread to fill the 400-μm pores. At 14 days, the number of cells increased slightly compared to that at 7 days. For TCs, Figure 3D shows low cell adhesion and proliferation. Even at 14 days, some of the larger pores were not completely filled. In addition, we found that the cells in the center of the pores are thinner than the cells attached to the fiber due to lack of support. With the completion of filling and further proliferation, the cells and secreted extracellular matrix can gradually fill the entire pore.

Figure 4 shows the results of the CCK-8 tests at 4, 7, 14, and 21 days after transplanting each group of cells on the scaffolds. First, every group showed an increase in measured values over time, indicating cell adhesion and proliferative activity. Among them, the 200 μm group showed the best result at 14 days for BMSCs. At 21 days, there was only little improvement over the 14 days for BMSCs and CCs. Furthermore, the 200 μm group showed the best result at 21 days, indicating that the environment of the 200-μm scaffold was best for BMSC adhesion and proliferation (Supplementary Figure 1). In the CC group, similar to the BMSC group, the results in each group improved continuously at 4, 7, and 14 days. However, unlike in the case of BMSCs, the 100 μm group showed similar results to the 200 μm group, considering that chondrocytes are small in size and more suitable for small pores. In the TC group, cell viability was lower than that of BMSCs and CCs at 4, 7, and 14 days but similar at 21 days compared to the other two cell types. At 21 days, the TCs showed best results in 300-μm pores, and the results were similar in 200-μm pores, which were only slightly lower than that in 300-μm pores. By contrast, 50 and 400 μm groups had the lowest values, suggesting that the environment of both 300- and 200-μm scaffolds was suitable for TC adhesion and proliferation. Taken together, the comparison of the three cell types demonstrated the different preferences of the cells in terms of pore size. Furthermore, all three cell types showed the lowest value in the 50 and 400 μm groups.

To investigate the most appropriate corresponding pore size for cartilage regeneration, BMSCs were implanted on different scaffolds. After cell adherence to the scaffold, the medium was replaced with a chondrogenic differentiation medium. Figure 5 shows the Sirius red and Safranine O-Fast Green staining stereoscopic images of the groups after chondrogenic differentiation on the scaffolds. In the images of the 50, 100, and 200 μm groups, the cells grew well with strong staining patterns. However, in the 300 and 400 μm groups, the cells did not fully grow and fill up all pores. Moreover, it was observed that collagen (Sirius red) and cartilage (Safranine O) in the 100 and 200 μm groups are more prominent.

The expression of COL II, COL I, AGC, and SOX9 after 21 days of stem cell induction is shown in Figure 6. Figure 6A shows the DAPI-labeled nuclei (blue), the phalloidin-labeled β-actin (green), the immunofluorescence for proteins (red), and the merged images. After further analysis, we found that the expression in the 100 and 200 μm groups was the best, especially compared to the 50 and 400 μm groups. This indicates that the pore size of chondrogenic differentiation had a clear range; too large or too small pores are not conducive to differentiation. The 100- and 200-μm pores were more suitable, the 50-μm pore was considered to be too small and oxygen and nutrients cannot be transported, while the 400-um pore is too large, such that the cells proliferate too much, and the differentiation ability was inferior.

Figures 7A–D show the mRNA expression of the chondrogenic differentiation marker genes (COL II, COL I, AGC, and SOX9) as determined by qRT-PCR using β-actin as an internal reference. The results were calculated by the 2^–ΔΔCt method. BMSCs were used as a reference, and the value was 1. We further clarified the effect of pore size on chondrogenic differentiation and further confirmed that too large or too small pores are not suitable for cartilage repair. In this part of the study, the 100 and 200 μm groups still showed good expression, while the 50 and 400 μm groups still had lower expression levels. These are consistent with the previous analysis, but, interestingly, in the previous section, the expression of COL II and AGC in the two groups of 100 and 200 μm showed the opposite results. Even if the difference is not obvious, we think that this small difference may be related to the sub-differentiation, such as superficial or mid-deep chondrocytes. Figures 7E–G show the biochemical evaluations after 21 days of stem cell induction. We examined the total weight of each group of scaffolds at 0 and 21 days and finally obtained the weight of the cellular components in the scaffold (Figure 7F). As with the previous studies, the 50 μm group had too small pores and low relative porosity. Oxygen and nutrients cannot be transported well, so cell adhesion and expression were the lowest. The 200 μm group was the highest and higher than the 300 and 400 μm groups. In addition, for the analysis of total collagen and type II collagen content, we found that the collagen content in the 50 μm group was not low, but the total amount was low. The collagen content gradually decreased with the increase of pores, being significantly decreased in the 400 μm group. Therefore, it may be that smaller pores are more capable of promoting collagen production. Thus, we believe that it is more appropriate to use scaffolds with different gradients for cartilage repair. It is more suitable to use smaller pores (such as 100 μm) in the surface layer and larger pores (such as 200 μm) in the middle and deep layers.

Although many studies have investigated the effect of scaffold pore size cells (Nehrer et al., 1997; Tanaka et al., 2010; Pok et al., 2013), due to the limitations of the manufacturing process and accuracy, the overall structure cannot be controlled entirely. With MEW technology, we can accurately prepare 50, 100, 200, 300, and 400-μm-pore-size scaffolds. The microstructure of the scaffold is suitable, and the fiber is smooth. At present, for MEW technology, some researchers have been able to control the fiber diameter to the hundred-nanometer level for stable printing. Although the material requirements are higher than other printing methods, it can provide higher printing accuracy (Hochleitner et al., 2015). Moreover, with the continuous development and application of 3D printing technology, it is vital to further study the effect of 3D printing scaffold microstructure on cells.

In our study, we first verified the impact of the difference in scaffold pore size on cell adhesion and proliferation and that the effect on different cells is different. For cell adhesion, we found in our study that pores that were too small (50 μm) may be more suitable for initial cell adhesion, such that this scaffold could provide more support for the cell. Even with very small pores, cells can easily pass through. During subsequent cell proliferation, cells on the small-pore scaffold were more likely to cross each other and fill the pores (Nuernberger et al., 2011). Therefore, in each group, the 50 μm pores were very easily filled with cells. Later, however, because microscopic pores do not provide enough space, cell proliferation is significantly limited in crowded environments, while slightly larger pores provide a broader environment for cells. However, very large pores take more time for the cells to grow and fill (400 μm) (Youssef et al., 2019). Moreover, it can be noticed that due to the lack of support from the scaffold, the cells in the center of the pores are lower than those on the fiber, and the pores are recessed, but with time, the recesses can be gradually filled.

In addition, the ability of different cells to adhere to the scaffold is different. As previously shown, CCs can climb up the scaffold and fill up the smaller pores more quickly. While it takes longer for larger pores, it still fills reasonably quickly (Bhardwaj et al., 2001; Griffon et al., 2006; Brennan et al., 2019). BMSCs have worse adhesion than CCs, with TCs being the worst. The observed phenomenon is the same as in the previous study, such that cells can easily pass through pores slightly larger than themselves, then slowly climb the scaffold at a faster proliferation rate (Tan et al., 2017). Conversely, studies on cell proliferation have shown that, as we have speculated before, different cells do have their optimal pore sizes, such as CCs were suitable for 100 and 200 μm, BMSCs for 200 μm, and TC for 200 and 300 μm. Thus, our findings provide novel insights for subsequent research, such that we can first explore the appropriate parameters before further repair of different tissues to enhance the repair effect (Salifu et al., 2017; Kankala et al., 2018). Moreover, although we cannot say that all the scaffolds will have the same trend, this similar trend should exist.

After studying the adhesion and proliferation of several cell types on MEW scaffolds with different pore sizes, we investigated the most appropriate corresponding pore size when BMSCs were induced into cartilage. Therefore, BMSCs were implanted on different scaffolds. Following cell adherence to the scaffold, the medium was replaced with a chondrogenic differentiation medium.

In our study, the results showed that the 100 and 200 μm groups were the most suitable for cell chondrogenic differentiation and gene expression. In immunofluorescence and qRT-PCR, the two groups exhibited apparent advantages over the other groups. This was significantly less in the 50 μm group, indicating that the smaller pores (50 μm) were not suitable for chondrogenic differentiation of stem cells.

Moreover, in a more in-depth study of chondrogenic differentiation of BMSCs, we found that microscopic pores were not conducive to cell growth and differentiation. For large pores, i.e., >300 μm, although they provided growth space, excessive cell proliferation also caused a decrease in differential expression. As such, completing tissue repair in a short period using tissue engineering scaffolds is desired, providing enough support to the cells without affecting their function (Gotterbarm et al., 2006; Brunger et al., 2014). Although there were no significant differences between the 100 and 200 μm groups in the study, we still have some interesting findings. In general, both groups could provide sufficient adhesion support for the cells, suitable growth space for the cells, and better performance. However, maybe due to differences in oxygen content or nutrient supply, COL II mRNA expression was higher in the 200 μm group.

Nevertheless, the total amount of COL II is higher in the 100 μm group. This indicates that pore size is also important for cell sub-differentiation. Therefore, in our study, we believe that the 200-μm-pore scaffold may be more suitable for the mid-layer cartilage scaffold, and a 100-μm-pore scaffold may be more suitable for the surface.

Three-dimensional printing technology can precisely control the geometry, mechanical structure, and spatial distribution of the manufacturing structure to mimic the normal structure (Jose et al., 2014; Thomas et al., 2014). Furthermore, different bioactive materials, cytokines, or cells can be loaded at different locations on a structurally controllable basis (Chang et al., 2015; Teimouri et al., 2015; Daly et al., 2016; Xu et al., 2018). Thus, accurate depictions and better replication of the complexity and heterogeneity of endogenous tissues and organs are expected with the construction of a controllable 3D system in the field of regenerative medicine. Therefore, the application of 3D printing technology in tissue engineering provides a new foundation for subsequent development (Lima et al., 2013).

For a tissue engineering scaffold, the gold standard should be its repair ability (Kang et al., 2016; Shi et al., 2017), for which many factors influence the final effect, such as the type of material, biological activity, printed porosity, and pore size, among others (Abdeen et al., 2014; Ryu et al., 2015; Zhu et al., 2018). However, further elucidation of the impact of these factors on the final repair is needed. To build large organizations, a better understanding of the combined effects of pore size on cell adhesion, differentiation, and secretion capacity is required (Chang et al., 2015; Daly et al., 2018; Prasopthum et al., 2018). Our results show that not all cells behave in the same manner on similar scaffold pores, and the difference in scaffold space is also different for the induction effect and protein expression. In the 3D printing process, the pore size can affect the porosity under other conditions unchanged. This correlation is very close, and because the porosity is not intuitive, the adjustment of the pore size is more direct for 3D printing. Thus, controlling the pore size of the scaffold may enhance tissue repair.

There are many shortcomings in this study. Due to the difficulty of standard setting and PCL material as a relatively inert synthetic material, it has yet to be tested in vivo. In addition, the experimental divisions (integer of 100, 200, and 300 μm) are not detailed enough; thus, a more detailed stratification may lead to better results. Considering the thickness of the articular cartilage of small animals in subsequent experiments, we only used 200-μm-thick scaffolds. Whether the increase of thickness has an effect on the results still needs further study. Besides this, the number of cell types and the direction of induction are somewhat insufficient. Therefore, to build a more detailed database, efforts by many researchers are needed.