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Combination of FACS and Homologous Recombination for the Generation of Stable and High-Expression Engineered Cell Lines

Traditionally, cell line generation requires several months and involves screening of over several hundred cell clones for high productivity before dozens are selected as candidate cell lines. Here, we have designed a new strategy for the generation of stable and high-expression cell lines by combining homologous recombination (HR) and fluorescence-activated cell sorting (FACS). High expression was indicated by the expression of secreted green fluorescent protein (SEGFP). Parental cell lines with the highest expression of SEGFP were then selected by FACS and identified by stability analysis. Consequently, HR vectors were constructed using the cassette for SEGFP as the HR region. After transfecting the HR vector, the cells with negative SEGFP expression were enriched by FACS. The complete exchange between SEGFP and target gene (TNFR-Fc) cassettes was demonstrated by DNA analysis. Compared with the traditional method, by integrating the cassette containing the gene of interest into the pre-selected site, the highest producing cells secreted a more than 8-fold higher titer of target protein. Hence, this new strategy can be applied to isolated stable cell lines with desirable expression of any gene of interest. The stable cell lines can rapidly produce proteins for researching protein structure and function and are even applicable in drug discovery.

Introduction
In recent years, the market for global biopharmaceuticals has widely expanded, and it is expected to exceed sales of US $166 billion by 2017 [1]. Major pharmaceutical products are recombinant proteins that are produced in cultivated mammalian cell lines, among which the Chinese hamster ovary (CHO) cell line is used to produce almost 70% of all recombinant protein therapeutics [2,3]. In the process of recombinant protein production, one of the critical steps is rapid selection of stable and high-expression cell lines for the gene of interest (GOI), which is a time-consuming and labor-intensive step [4]. To generate cell lines for the production of target proteins, the traditional strategy involves transfection of the target gene for random integration into genomic DNA by homologous recombination (HR). The titer of the target protein is then analyzed among a large number of cell clones to select high-expression cell clones. Using this method, more than 80% of cell clones express the GOI at a very low level. Even in high-expression cell clones, GOI expression needs to be increased by several rounds of amplification. Lastly, single cell clones can be isolated by subcloning [5,6]. Furthermore, the selected cell clones have some limitations, such as instability and/ or slow cell growth [7]. The most important step of this procedure is integration of the GOI into a stable and high-expression site in the genomic DNA, which enables high and continuous expression of the GOI. Therefore, in modern biopharmaceutical technology, different strategies have been developed to increase the screening throughput of cell clones and/or raise GOI expression directly.
More than 100 million cells are used to establish one cell line for recombinant protein production [6]. To obtain more cell clones, many more cells need to be analyzed and rapidly selected by highthroughput screening. Fluorescence-activated cell sorting (FACS) is a widely used method for rapid analysis of a large number of cells [8]. There are several strategies that can be applied to this technology: 1) green fluorescent protein (GFP) as a reporter gene for selection of GOI high-expression cells [9]; 2) immunostaining using an antibody or Fc-fusion protein and sorting the highly fluorescent cells that indicate high-expression cells [10]; 3) selection of a new host cell line from a large number of cells to generate the GOI high-expression cell line [11,12]. On the other hand, cell clones can be analyzed by flow cytometry at the early stage to determine their stability [13].
Very different strategies have been developed to increase GOI expression, including insertion of an increased expression element or using a new promoter to increase transcription of the GOI [14,15]. These strategies include using STAR/MARs/UCOE elements to reduce gene silencing induced by epigenetic effects [16][17][18], selection of cell lines containing a hotspot region for high expression, as indicated by a reporter gene, and integration of the GOI into these regions using Cre-LoxP and/or Flp-In systems [19,20]. All of these strategies would save time and reduce costs to obtain high-expression cell lines.
In this study, we report a new strategy for establishment of a GOI high-expression cell line. By combining HR and FACS, our strategy was designed to enrich and collect the gene-replaced cells that exchanged a secreted GFP (SEGFP) cassette with the GOI cassette at a hotspot in the genome. Compared with the traditional method, our results revealed that the titer of GOI-encoded protein increased approximately 8-fold by insertion of the GOI into the pre-selected site. The GOI-engineered cell lines inherited the stable cell growth and protein expression of the parental cell line. Therefore, to construct stable and high-expression cell lines, only a small number of cell clones needs to be generated and analyzed using our strategy, which saves time and significantly reduces costs. Considering these advantages, combining HR and FACS provides a new and rapid strategy for the generation of stable and highexpression cell lines.

Materials and Methods
Plasmids SEGFP contains a signal peptide from the human IgG kappa chain in the N-terminal. The expression cassette was composed of a CMV promoter, SEGFP coding region, and BGH polyA sequence.
To construct a TNFR-Fc expression control plasmid, a DNA fragment of the human EF1a promoter and TNFR-Fc gene was amplified by PCR from the proprietary TNFR-Fc expression vector. The amplified DNA fragment was digested with PvuI/ EcoRI, and then inserted into a pBudCE4.1 plasmid (Invitrogen, USA) to replace the CMV promoter.
All HR plasmids were constructed from the control vector. For HR plasmid A, the full-length CMV sequence and 400 bp SEGFP N-terminal region were amplified, digested by PvuI, and then inserted upstream of the human EF1a promoter. The 300 bp SEGFP C-terminal sequence and BGH polyA signal sequence were cloned and inserted downstream of the zeocin-resistance gene after digestion by NheI.
HR plasmid B was obtained by ligating the NheI-digested SEGFP c-polyA-Neo p fragment into digested plasmid A. The SEGFP c-polyA-Neo p fragment contained the SEGFP C-terminal sequence region (300 bp) to the partial Neo gene.
HR plasmid C was also derived from plasmid A, in which the upstream HR element was replaced by a partial CMV promoter (CMV p; 200 bp).

Cell Culture
CHO-S cells were purchased from Invitrogen and cultivated in CD-CHO medium (Gibco/Invitrogen, USA) containing 8 mM Lglutamine and HT supplement (100 mM hypoxanthine and 16 mM thymidine). Host cells and cell clones were cultured in 100 ml shaking flasks containing 20 ml medium at 37uC in a 5% CO 2 incubator. Cell concentration and viability were measured by trypan blue exclusion using a hemocytometer. Routine passaging was performed at a frequency of three times per week. SEGFP cell clones were cultured under the same condition with growth medium containing 0.6 mg/ml G418. Recombinant cell pools and clones were cultured in medium containing 0.1 mg/ml zeocin.
Population doubling time (PDT) was calculated from the culture time versus a semi-log plot of the viable cell concentration in the exponential growth phase by the following formula: where x 1 and x 2 are the viable cell concentrations at time points t 1 and t 2 .
Integral cell area (ICA) was calculated by the following equation: where N and N 0 represent the final and initial viable cell numbers, respectively, while t represents the days in culture.

Genes Transfer
Electroporation. CHO-S cells were seeded at 24 h before transfection. A total of 5610 6 cells and 10 mg DNA were used for transfection at 350 V for 30 ms in a Gene-Pulse (Bio-Rad, USA). The transfected cells were resuspended in CD-CHO medium and cultured in a shaking flask with 10 ml of culture medium. For selection of cells as a control, at 1 day post-transfection, the electroporated cells were plated in 24-well plates with culture medium containing 0.1 mg/ml zeocin.
Transfection of HR plasmids. As described above, at 1 day prior to transfection, SEGFP-transfected cell clones were seeded in a shaking flask. A total of 5610 6 logarithmically growing cells were electroporated with HR plasmids (10 mg). The transfected cells were recovered in a shaking flask containing growth medium. At 72 h after transfection, transfected cells were collected for SEGFPnegative cell sorting.

Measurement of SEGFP and TNFR-Fc
Fluorescence Intensity (FI) measurement. Culture medium of each SEGFP cell clone was collected and centrifuged at 13,000 g for 10 min to remove cell debris. The medium supernatant was then transferred to a fresh 1.5-ml tube. A total of 200 ml medium supernatant was aliquoted into 96-well plates. Fresh medium was used as a blank control. The plates were then scanned on a Cytofluorescent Synergy TM 2 Fluorescence Measurement System (Biotek, USA) with filter sets covering the GFP excitation and emission wavelengths (excitation: 485620 nm; emission: 530625 nm).
The quantitative FI (QFI) was calculated with the FI by the following equation:

QFI~FI=ICA
where FI is the total FI in the culture supernatant as determined by spectrofluorometric measurement.
Enzyme-linked immunosorbent assay. To determine TNFR-Fc titers, conditioned medium was analyzed by the sandwich enzyme-linked immunosorbent assay method. Essentially, 96-well plates (Constar/Corning, USA) were pre-coated with a goat anti-human IgG primary antibody (CW bio, China) at 37uC overnight. Following a series of wash steps, the wells were blocked with bovine serum albumin in PBS and Tween 20. Bound recombinant antibodies were detected using TMB substrate (Invitrogen, USA) following incubation of each well with a mouse anti-human IgG HRP-conjugated secondary antibody (CW bio, China). Following 30 mins of incubation at room temperature, the TMB substrate reaction was terminated with 2 M H 2 SO 4 prior to reading at 450 nm using a microplate reader (Thermo, USA). Data were collected and analyzed using GraphPad Prism 5. Quantification was based on a dilution series to obtain a standard curve of known TNFR-Fc concentrations.
Western blotting. SEGFP and TNFR-Fc proteins in culture supernatants were analyzed by western blot. An aliquot of growth medium was separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis. The proteins were then transferred onto a polyvinyl difluoride membrane, and the presence of SEGFP or TNFR-Fc was demonstrated using anti-GFP (CW bio, China) and anti-human IgG antibodies, respectively, following by DAB staining (Tiangen, China).

Flow Cytometry
Cells prepared as described above were resuspended in cold PBS at approximately 1610 6 viable cells/ml, followed by analysis using a FACSCalibur flow cytometer (BD Biosciences, USA). Auto fluorescence of untransfected cells was detected as a control. Data acquisition was carried out by analyzing 10,000 events/sample using Cell Quest Software (BD Biosciences, USA).

FACS
At 3-4 days after transfection, exponentially growing cells were harvested and resuspended in culture medium with 1% penicillin/ streptomycin at approximately 5610 6 viable cells/ml. The collected cells were then sorted on a FACStar PLUS. Live cells were gated based on forward/side scatter and events in the top 3-4% of GFP fluorescence were sorted. Seven days later, a second round of cell sorting was performed to collect cells in the top 1-2% of GFP fluorescence. The sorted SEGFP high-expression cells were cultured in medium containing 0.6 mg/ml G418.
For HR cells, fluorescently gated cells in the lowest 5-6% (first round) and 2-3% (second round) were sorted. Both groups of sorted cells were cultured in growth medium with 0.1 mg/ml zeocin. SEGFP high-expression cells and HR cells from the second round of sorting were expanded and subcloned at a concentration of 1 cell/well in 96-wellplates.

DNA Analysis and Copy Number Measurement
Genomic DNA was harvested from cells with a Genomes DNA extraction Kit (TaKaRa, Japan). PCR was performed with HS Primer STAR polymerase (TaKaRa, Japan) according to the manufacturer's instructions using a MJ (Bio-Rad, USA) as follows: pre-denaturation at 94uC for 3 min, followed by 35 cycles of denaturation at 94uC for 30 s, annealing at 65uC for 1 min, and elongation at 72uC for 2 min. The primers used were as follows: CMV-S: 59 GTACATTTATATTGGCTCATGTCCAACAT-TACCGC 39 SEGFP-A:59 GGCGGACTGGGTGCTCAGGTAGTGGTT 39 TFNR-A:59 AAGATTTGGGCTCAACGCTACCTCCACC 39 DNA was extracted from cells using an All Prep DNA Mini kit (Qiagen, Germany), and then applied to qPCR (SYBRH Premix Ex Taq TM kit; TaKaRa, Japan) to determine the copy number of the SEGFP gene using the following primers: forward: 59 TGCTTCAGCCGCTACCC 39 and reverse: 59 TCACCTT-GATGCCGTTCTT 39. PCR was performed on an Opticon monitor II (Bio-Rad, USA) at 95uC for 30 s, followed by 45 cycles of 95uC for 5 s and 60uC for 30 s. The linearized SEGFP vector was used to generate a standard curve, and copy numbers were determined using the absolute quantitative method.

Results
In the pharmaceutical industry, establishing a stable and highexpression cell line is very labor-intensive and highly inefficient. To improve the traditional method, we considered generation of a high-expression cell line based on HR. As shown in Fig. 1, to obtain the GOI high-expression cell line, high-expression parental cells were first selected using SEGFP as a marker. A high expression level of SEGFP indicated that its insertion site was a hotspot region for exogenous protein expression. In addition, the SEGFP expression cassette was used as the homologous sequence to construct the HR vector. In the HR vector, the CMV promoter-SEGFP N-terminal sequence and SEGFP C-terminal-polyA signal sequence were inserted into the upstream and downstream sequence of the GOI expression cassette, respectively. According to this design, the GOI would be inserted in the preselected site by replacing the SEGFP expression cassette with the HR region to easily establish the GOI high-expression cell line. This strategy included the following key points: selection of SEGFP high-expression cell lines, construction and functional analysis of HR vectors, collection of SEGFP-negative cells, and measurement of GOI expression in replaced cells (SEGFPnegative cells).

SEGFP Parental Cell Line Generation
The most important point of this strategy is selection of a parental cell line with high expression of the reporter gene. SEGFP was used as a marker to select the high-expression cells by two rounds of FACS. Cells with the highest expression of SEGFP (top 3-4%) were sorted on day 4 after transfection. After 7 days, the top 1-2% of cells with high expression of SEGFP were sorted again ( Fig. 2A). After the sorting process, almost all sorted cells were SEGFP positive. The percentage of SEGFP high-expression cells was increased from 81.8% (first-sorted cells) to 99.1% (final sorted cells).
To select a SEGFP high-expression parental cell line, the cells from the second round of FACS were individually cultured in 96well plates. After comparing the expression of SEGFP, the top 10 clones were chosen from among 285 clones to analyze the stability of cell growth and SEGFP expression. Fig. 2B shows the relationship of quantitative fluorescence intensity (QFI), Population doubling time (PDT) and total FI among these clones. The average PDT of the SEGFP clones ranged from 19 to 22 h, except for clones 6A5 (29.4 h), 20E2 (29.02 h) and 2-1 (24.83 h). Consistent with their PDTs, the average QFI was higher in the three clones than that in other clones. Average QFI values among these clones had a more than 2-fold divergence from 4.3 to 9.58610 23 FI/c/d. However, the discrepancy was only 25% between the highest and lowest FI. Furthermore, all SEGFP clones were single-positive clones as determined by flow cytometry. The coefficients of variations (CV) in different clones ranged from 4.3 to 7.5 (data not shown).
Combined with QFI and total FI, four SEGFP clones (gray circles in Fig. 2B: 6A5, 2-1, 1-14, and 1D1) were selected to carry out the HR experiment. The characteristic parameters of the four SEGFP clones are summarized in Table 1, including PDT, QFI, percentage of positive cells (positive %), CV, and copy number. The four clones showed two different sets of characteristics: 6A5 and 2-1 cell clones grew slower with a higher QFI and lower uniformity, whereas 1-14 and 1D1 cell clones grew faster with a lower QFI and higher uniformity. Moreover, the stability of these clones in terms of cell growth and SEGFP expression was analyzed by the average PDT (PDT AVG ) and average QFI (QFI AVG ), respectively. The PDT AVG of these clones changed smoothly, indicating that these clones had stable growth over multiple passages (Fig. 2C). The variation of QFI AVG revealed that the SEGFP expression levels in 2-1, 1-14 and 1D1 cell clones were stable at the start of subculture. After adaption to the shaking flask, SEGFP was expressed stably in the 6A5 cell clone beginning from passage 8 (Fig. 2D).

Construction of HR Vectors
In parental cells, the SEGFP cassette from the CMV promoter to the polyA signal sequence had to be inserted into the genomic DNA. Therefore, HR vectors derived from the TNFR-Fc expression vector were constructed by inserting the homologous sequence of the SEGFP gene into the upstream and downstream sequence of the GOI (Fig. 3A).
Three different vectors were constructed for HR. Plasmid A contained the full-length CMV promoter and a partial sequence of the SEGFP N-terminal (1100 bp) in the upstream and a partial sequence of the SEGFP C-terminal plus polyA region (700 bp) in the downstream. The downstream element of plasmid B was increased to 1700 bp by adding a partial Neo gene sequence behind the polyA region, and the upstream sequence had no changes from plasmid A. In plasmid C, the size of the upstream element was decreased to 200 bp, which only contained a partial CMV promoter, and the downstream element remained unchanged from plasmid A.
First, the plasmid construction should not inhibit GOI expression. To detect the expression of the GOI (TNFR-Fc) from the different HR vectors, we transfected these plasmids into CHO cells and used the TNFR-Fc expression vector as a control. After electroporation, the cell growth of transfected cells remained the same. However, compared with the TNFR-Fc titer produced by the control vector (6 mg/L), only plasmid C could express a higher TFNR-Fc titer (10 mg/L). The Qp of plasmids A and B was also much lower than that of the control (Fig. 3B). Thus, plasmid C was used for HR.

Selection of SEGFP-negative cells by FACS and
comparison of sorted cell pools. According to the design, the SEGFP gene would be replaced by the GOI expression cassette after transfection of HR plasmids. Thus, we collected SEGFP-negative cells, even though not all SEGFP-negative cells represented replaced cells. The flow chart in Fig. 4A shows the two-round selection procedures to obtain SEGFP-negative cells by FACS. In the first round of sorting, cells in the lowest 5-6% of SEGFP expression were selected and designated as the ''-S1'' cell pool. After recovery in 24-well plates, the 2-3% SEGFP lowestexpression cells were then sorted and designated as the ''-S2'' cell pool.
The cell pools were expanded and analyzed. In all four cell clones, the SEGFP lower-expression cells were enriched after the two rounds of sorting (Fig. 4B). However, the cell pools selected from different parental cell lines had a different pattern of SEGFP expression. For example, the first cell pools (1D1-S1 and 1-14-S1) derived from 1D1 and 1-14 cells consisted of the lower 5-6% SEGFP-negative cells and almost all cells were SEGFP negative in the second cell pools (1D1-S2 and 1-14-S2). In contrast, SEGFP expression in 6A5-S2 and 2-1-S2cell pools was also decreased significantly, but SEGFP-negative cells were not enriched by FACS. The cell size and protein content were also compared between sorted cells and parental cells by flow cytometry (Fig. S1). No significant variations were observed among sorted cell pools and parental cell lines, although the four cell clones had different cell sizes and protein contents.
In addition to comparison of SEGFP expression, cell growth and TNFR-Fc expression were compared in sorted cell pools ( Fig. 4C and 4D). The average PDTs of 2-1-S2, 1-14-S2 and 1D1-S2 cells were unchanged, compared with that of the first-sorted cell pools. All of the cell pools had PDTs of approximately 20 h, suggesting that the cells selected by FACS could grow very well. However, the 6A5-S2 cell pool had a significantly longer PDT (39.84 h) than that of the 6A5-S1 cell pool (26.67 h) (P = 0.026). Following FACS, the average Qp of TNFR-Fc was increased compared with that of the corresponding cell pools derived from different cell clones. The Qp of 1D1-S2 and 1-14-S2 cell pools was 2.60 and 0.66 pg/cell/day, respectively. Although 2-1-S2 cells were SEGFP negative, the Qp of TNFR-Fc in the cell pool increased to 3.96 pg/cell/day. These results demonstrated that SEGFP-negative or lower-expression cells can be collected and enriched by FACS, the SEGFP-negative cells expressed TNFR-Fc, and the Qp of TFNR-Fc increased along with the increase of SEGFP-negative cells.
Site-integration analysis of HR cells. In accordance with the design, SEGFP-replaced cells would be SEGFP-negative cells, but this assumption was incorrect. Therefore, even though 1D1-S2 and 1-14-S2 cell pools were almost completely composed of GFPnegative cells, as detected by flow cytometry, we also determined whether these cells were replaced cells by another detection method. Consequently, medium supernatants of all four cell pools were collected and analyzed by western blotting. As shown in Fig. 5A, only SEGFP was expressed in the four SEGFP cell clones, and the expression level was consistent with their FI. Moreover, only one band for SEGFP was detected in the medium supernatants of the four sorted cell pools, suggesting that SEGFP was not replaced by the different expression cassettes by HR. Consistent with the flow cytometry results, the SEGFP expression level decreased in all sorted cell pools. Notably, in the 1D1-S2 cell pool, no observed SEGFP band suggested that the SEGFP expression cassette was replaced completely. On the other hand, in all sorted cell pools, TNFR-Fc was expressed at different levels.
To confirm whether HR was complete, we isolated the genomic DNA of SEGFP cell clones and the corresponding sorted cell pools. The reporter gene and GOI were then detected by PCR. As shown in Fig. 5B, we designed three primers for gene amplification. The same upstream primer was used to amplify SEGFP and GOI in the respective DNA samples. It was designed to match the upstream CMV promoter in the SEGFP vector other than the HR vector. Thus, in SEGFP cell clones, the DNA fragment (1700 bp) from the CMV promoter to the C-terminal of SEGFP was amplified using a CMV-S and SEGFP-A primer pairs; without TNFR expression cassette, no band was detected in these genomic DNA using CMV and TNFR-A primers. However, after HR and cell sorting, the DNA band was shifted in site-integrated cells, in which the DNA fragment of CMV-TNFR (2500 bp) was amplified using CMV and TNFR-A primers. Because of mismatching with the sense primer of the CMV promoter, no band was amplified from the HR vector.
In accordance with the flow cytometry and western blot results, the corresponding fragments were amplified from SEGFP cell clones using the CMV-S and SEGFP-A primer pair (Fig. 5C). Different patterns were revealed in different sorted cell pools. The shifted band (2500 bp) was detected only in 1D1 and 2-1 cell pools using CMV and TNFR-A primers, suggesting that the GOI cassette was inserted into the pre-set site of genomic DNA of 1D1 and 2-1 cell clones. However, no specific bands were observed in 6A5-S2 and 1-14-S2 cell pools, indicating that the GOI cassette did not replace the SEGFP site exactly. On the other hand, SEGFP was amplified in all sorted cell pools using CMV-S and SEGFP-A. Some SEGFP-positive cells in 6A5-S2 and 2-1-S2 cell pools may be the reason for this result. Detection of SEGFP in the 1-14-S2 cell pool indicated different HR patterns. The unexpected results of the 1D1-S2 cell pool may have been caused by a few cells containing the SEGFP gene that was not expressed.

Increased GOI Expression by Integration in Pre-selected Sites
Higher GOI expression in the SEGFP-negative cell pool than that in the SEGFP-positive cell pool. To compare the expression level of the GOI with site-specific or random integration, SEGFP-negative cells (GFP 2 cells) and SEGFPpositive cells (GFP + cells) were separated by FACS (Fig. 6A). As shown above, the GOI in the 1D1-S2 cell pool was integrated into the genomic DNA at a pre-selected site. Thus, the sorted GFP 2 and GFP + cells represented site-integrated and randomly integrated cells, respectively. Using the 1D1-S1 cell pool as a control, the cell growth and GOI titer of sorted cell pools were compared. The average PDT of the cell pools was approximately 20 h (Fig. 6B), indicating that cell growth had not been inhibited by SEGFP and/ or GOI expression. However, the averaged Qp of the GOI in SEGFP-negative cells was significantly higher than that in SEGFP + cells and the 1D1-S1 cell pool (Fig. 6C). These results suggested that the GOI integrated into the high-expression site will produce more target protein than that of the randomly integrated GOI.
GOI expression increases in cell pools and clones generated by HR. To verify that the cells with the GOI integrated into the pre-selected site (1D1-S2 cell pool) can express a higher amount of target protein, we transfected HR plasmid C into CHO cells and generateda cell pool by traditional drug selection (CHO cell pool). Cell growth and Qp of both cell pools were compared during subculture in shaking flasks. The PDTs of  the cell pools were not significantly different, although the PDT of the 1D1-S2 cell pool (18.1 h) was slightly lower than that of the CHO cell pool (19.3 h) (Fig. 7A).On the other hand, as shown in Fig. 7B, the Qp value of the 1D1-S2 cell pool was approximately 2.6 pg/cell/day, which was more than 5-fold higher than that of the CHO cell pool (0.4 pg/cell/day).
Cell pools generated by the traditional method were composed of a large number of heterogeneous cells with different expression levels of the GOI. Therefore, the advantage of HR could not be demonstrated by comparing cell pools. Forty-six and 42 cell clones were then selected by subcloning CHO and 1D1-S2 cell pools, respectively. According to high GOI expression, seven CHO cells clones and eight 1D1-S2 cell clones were expanded. As shown in Fig. 7C, the GOI in 1D1-S2 cell clones was expressed significantly higher than that in cell clones derived from the CHO cell pool.
The Qp values of 1D1-S2 cell clones was 2.8-4.1 pg/cell/day, which was slightly higher than that of the 1D1-S2 cell pool. The PDTs were equivalent among cell clones from different cell pools (Fig. 7D). TNFR-Fc titers of cell clones selected from the 1D1-S2 cell pool were much higher than those of CHO cell clones. Notably, the TNFR-Fc titer of the best 1D1-S2 cell clone was 38.6 mg/L, which was approximately 8-fold higher than that of the best CHO cell clone (Fig. 7E). On the other hand, among cell clones, only the GOI gene was amplified by CMV and TNFR-A primers, and no SEGFP gene fragment was detected by CMV and SEGFP-A primers in All 8 single cell clones (Fig. 7F). The band shift showed an exact gene replacement between the reporter gene and GOI. In conclusion, cell clones from the 1D1-S2 cell pool showed higher GOI expression, and they were composed by cells without the SEGFP gene.

Discussion
In this study, we present an alternative approach for the generation of high-expression engineered cell lines. Stable cell lines with significantly increased expression of TNFR-Fc were established by combining HR and FACS (Fig. 7E). Furthermore, the cell pool derived from our strategies can express a GOI at a high level, which can be used to produce recombinant protein. This strategy also provides time and cost advantages over traditional processes.
In the biopharmaceutical industry, stability and high-expression, which result from GOI insertion into a genomic hotspot [5], are two critical aspects for engineered cell lines. Our strategy fist established a parental cell line with genomic hotspots indicated by a reporter gene. HR was then used to insert the GOI into the preselected site to generate the engineered cell line. Some guiding principles can be summarized from our results of parental cell line generation and the following HR.
In general, the highest expression of the reporter gene in the parental cell line was acquired in batch culture. It has been reported that an increase in specific productivity directly correlates with the cell size and protein content [21,22]. Our flow cytometry data also showed that cell clones with a higher cell content and larger cell size had a higher QFI (data not shown). However, the titer of a target protein in batch culture is determined by cell density as well as Qp. Porter et al. reported that clones with the highest GOI expression are those with a higher Qp and the highest visual cell density rather than those with the highest Qp [23]. There was only a 15% difference of total FI in our four clones, among which there was an approximate 2-fold difference of QFI. Thus, selection of the parental cells should balance the Qp and cell growth. On the other hand, HR results suggested that the cell clones, such as 1D1 and 1-14, with a higher QFI, high cell density, and shorter PDT may have an advantage for gene replacement.
The replaced cells displayed a consistent cell size and protein content compared with that of their parental cells (Fig. S1). Therefore, the stability of the parental cells was important for the stability of the engineered cell lines, including the stability of cell growth and protein production. These stable cell clones were all single cell clones with significant homogeneity of the cell population, resulting in little difference of antibody or Fc-fusion protein expression in the cell population [13]. In parental cells, the homogeneity of SEGFP expression was easily determined by flow cytometry. 1D1 and 1-14 cell clones had better homogeneity than that of the other two clones (Table 1). On the other hand, the 6A5 cell clone had more heterogeneous cells with significant instability of cell growth and SEGFP expression. Moreover, cell clones of GOI, derived from 1D1-S2 pool, exhibited similar characters, such as Qp, PDT and HR patterns (Fig. 7D-F). Thus, homogeneity is an important character for selection of parental cells to generate stable gene-replaced cells for both cell growth and GOI expression. Following these principles, much more SEGFP cell clones were selected in the next round. The cell clones of GOI with higher-expression and extract HR replacement were generated by using those SEGFP parental cell clones, similar with 1D1 (data not shown).
Copy number of the reporter gene in parental cells is also an important factor. In theory, a single copy of the reporter gene is required for gene replacement with the GOI [19]. More than one copy of the reporter gene will significantly reduce the HR efficiency [24]. However, our results indicated that not only a single copy of the reporter gene can be replaced by HR; it has also been reported that high-expression cell lines can be established by HR of a parental cell line containing two copies of the reporter gene [25]. Therefore, to increase the efficiency of HR and obtain more replaced cells, the parental cells should contain the lowest possible copy number of the reporter gene.
In addition to establishment of parental cell lines, construction of the HR vector is also very important. HR plasmid C, which expressed the GOI as well as the control plasmid (Fig. 3B), was used to insert the GOI into the hotspot. Moreover, this construction can increase the GOI expression level when the plasmid randomly integrates into other genomic sites in addition to the pre-selected site. Our results indicated that only a 200 bp homologous region could guide HR between the GOI and reporter gene. Even without recombinase, the 34 bp minimal Flp recognition target (FRT site) can increase the probability of HR (data not shown). One of the reasons is that electroporation can increase the efficiency of DNA importation into the nucleus, which improves the probability of HR [26,27].
Hasty et al. reported that only 472 bp of homology is used as efficiently as 1.2 kb in the formation and resolution of cross over junctions [28]. However, to obtain HR cell clones, the size of the homologous region has to be more than 1000 bp. Following these principles, HR plasmids A and B were constructed simultaneously with a longer homologous region. However, all constructions can indicate, respectively,similar DNA structure after HR. GOI expression was very low using transient transfection of plasmids A and B, suggested that the GOI may be expressed at a very low level in genomic DNA. To determine whether GOI expression was inhibited by transcription of the SEGFP N-terminal under the full-length CMV promoter, HR plasmids A and B were also improved by inserting the TK polyA sequence following the SEGFP N-terminal sequence. However, the decrease of GOI expression did not recover using the improved plasmid (data not shown). In our study, the efficiency of HR was very low, even though we increased the efficiency using the homologous region. However, using SEGFP as the reporter gene, HR cells should be SEGFP-negative cells that could be easily enriched by FACS. As expected, almost all SEGFP-negative cells in 1-14-S2 and 1D1-S2 cell pools were selected by the two-round sorting process (Fig. 4B). On the other hand, complete HR could not be ensured by SEGFP-negative expression. Integration of the GOI expression cassette in the pre-selected site should be determined by expression of the GOI and analysis of genomic DNA. Analysis of 1-14-S2 and 1D1-S2 cell pools demonstrated that the HR process could result in different replacement patterns. Furthermore, with differences among 1D1-S2 cell clones, the SEGFP gene was also detected in 1-14-S2 cell clones (data not shown). In summary, SEGFPnegative cells can be enriched by FACS, but GOI expression and DNA analysis are needed for exact determination of replaced cells.
Different aspects of the engineered cell line establishment need to be improved, although high-expression cell lines can be selected with our strategy. First, it has been reported that the S/MARs region can increase reporter gene expression, and increase expression of a GOI inserted by RMCE. However, the limitations of gene expression with a single copy are unknown. In addition, the expression level of the reporter gene in the parental cell line should be as high as possible. However, accumulation of the reporter gene may affect cell growth. Our use of SEGFP would decrease the accumulation of intracellular GFP, and then reduce the inhibiting effect of GFP on cell clone selection and cell growth. However, SEGFP would also be expressed in cells, which may affect selection of high-expression cells. Thus, to indicate the best parental cell line with a high-expression site, we need to improve the expression cassette of the reporter gene or construct a more suitable reporter gene.


Activation of intervertebral disc cells by co-culture with notochordal cells, conditioned medium and hypoxia

Background Notochordal cells (NC) remain in the focus of research for regenerative therapy for the degenerated intervertebral disc (IVD) due to their progenitor status. Recent findings suggested their regenerative action on more mature disc cells, presumably by the secretion of specific factors, which has been described as notochordal cell conditioned medium (NCCM). The aim of this study was to determine NC culture conditions (2D/3D, fetal calf serum, oxygen level) that lead to significant IVD cell activation in an indirect co-culture system under normoxia and hypoxia (2% oxygen). Methods Porcine NC was kept in 2D monolayer and in 3D alginate bead culture to identify a suitable culture system for these cells. To test stimulating effects of NC, co-cultures of NC and bovine derived coccygeal IVD cells were conducted in a 1:1 ratio with no direct cell contact between NC and bovine nucleus pulposus cell (NPC) or annulus fibrosus cells (AFC) in 3D alginate beads under normoxia and hypoxia (2%) for 7 and 14 days. As a positive control, NPC and AFC were stimulated with NC-derived conditioned medium (NCCM). Cell activity, glycosaminoglycan (GAG) content, DNA content and relative gene expression was measured. Mass spectrometry analysis of the NCCM was conducted. Results We provide evidence by flow cytometry that monolayer culture is not favorable for NC culture with respect to maintaining NC phenotype. In 3D alginate culture, NC activated NPC either in indirect co-culture or by addition of NCCM as indicated by the gene expression ratio of aggrecan/collagen type 2. This effect was strongest with 10% fetal calf serum and under hypoxia. Conversely, AFC seemed unresponsive to co-culture with pNC or to the NCCM. Further, the results showed that hypoxia led to decelerated metabolic activity, but did not lead to a significant change in the GAG/DNA ratio. Mass spectrometry identified connective tissue growth factor (CTGF, syn. CCN2) in the NCCM. Conclusions Our results confirm the requirement to culture NC in 3D to best maintain their phenotype, preferentially in hypoxia and with the supplementation of FCS in the culture media. Despite these advancements, the ideal culture condition remains to be identified. Electronic supplementary material The online version of this article (doi:10.1186/1471-2474-15-422) contains supplementary material, which is available to authorized users.

Background
The notochord is a key structure in early embryogenesis in all chordates and is populated with relatively big and vacuole enriched notochordal cells (NC) [1,2]. The cells form the center of the intervertebral disc (IVD) in the fetus and the adult [3,4]. These cells are either present throughout the lifetime at various ratios across the vertebrate phylogenetic tree with respect to other IVD cells, or they are reduced to a minimum population after adolescence [5,6]. In humans, it has been found that they disappear early in childhood [7]. It has been described that in different mammalian groups, these cells co-exist with nucleus pulposus cells (NPC) at different ratios [5,7]. For instance, in cattle, goats, and sheep, these cells disappear early in lifetime [3,7], whereas in rodents (rats and mice) and rabbits, a high number of NC is maintained throughout their lifetime. In dog breeds, both IVD types are present. In the so-called non-chondrodystrophic dogs (e.g. larger dogs, such as greyhounds) in which low back pain is less common, a high ratio of NC is present. Conversely, in chondrodystrophic dog breeds (e.g. beagle, dachshund), few NC are present, and these breeds frequently suffer from disc degeneration and low back pain. Over the past decade, the fate of NC has been a matter of debate. It was hypothesized that these cells either disappear by apoptosis or that they are differentiated into an IVD chondrocytelike cell phenotype [8,9]. On this matter, recent transcriptome analyses of the possible origin of these cells have pointed into the direction of precursor origin and that these two cell populationsalthough very different in size and granularitywere not so different in their phenotypic profile [10,11]. The open question remains about the mesodermal or ectodermal origin of these cells [12][13][14]. However, recently, "noto," a highly conserved homeobox gene restricted to the notochord, where it regulates node morphogenesis, has been tracked by a novel notochord-specific Cre mouse. These gene-tracking data, during embryogenesis, clearly demonstrated that the notochord indeed gives rise to the NP and later, to the smaller chondrocyte-like disc cells, which confirms the close relationship of the NC and the smaller NPC [15][16][17]. However, the interaction between these morphologically different cell types remains obscure, hence leading to cross-species co-cultures investigations [13,14,18] and conditioned medium experiments [13,14,[19][20][21][22][23]. In a dog model, it was found that NC secrete connective tissue growth factor (CTGF, syn. CCN2) and can protect NPC from undergoing apoptosis [24,25] by inhibiting the caspase pathway. It was furthermore demonstrated that NC are highly sensitive to the culture conditions, i.e., to media composition and oxygen content [26]. More importantly, NC or their cultured mediathe so-called notochord-derived conditioned medium (NCCM)demonstrated regenerative effects by inducing the glycosaminoglycan per DNA (GAG/DNA) ratio [13,18,21]. These cells, or, possibly, merely their released factors, are of high interest for regenerative medicine to re-populate degenerated IVDs or to stimulate native disc cells. Our aim was to test whether NC can activate NPC in vitro and which initial culture conditions may be favorable to activate more differentiated disc cells. The experimental set-up (explained below) is shown in Figure 1. Our first step was to revisit the question for suitable culture conditions for these cells. Previous cell culture studies expanded NC in 2D monolayer [14] and also, more recently, in 3D [26,27]. It has been noted that NC are highly sensitive to mechanical loading and changes to the nutritional environment [26][27][28][29]. What is currently missing is a direct proof that 2D culture conditions are indeed unfavorable for NC and that the activation capacity of NC on other cells is dependent on the nutritional state of the NC in culture. Thus, we wanted to demonstrate the effect of 2D monolayer culture versus 3D cell culture in alginate. Furthermore, we investigated the hypothesis that NC are sensitive to culture conditions [28] and that their activation capacity on disc cells in indirect co-culture experiments can be stimulated by exposure to fetal calf serum (FCS). As a positive control, we included the previously described application of conditioned medium (NCCM) collected from NC that has been demonstrated to stimulate smaller NPC [21,22,27,30]. However, it is unknown whether annulus fibrous cells (AFC) can be activated by either coculture or exposure to NCCM. We then also replicated the experiments in normoxic (~21%) and hypoxic (~2%) conditions to study the effect of the physiological oxygen environment as it was also reported in the literature that NC should be preferentially cultured in hypoxia for a better aligned extracellular matrix [20,26] and a more authentic phenotype as defined by the production of aggrecan with respect to collagen type 2, which was found to be around 27:1 in human tissue [31] or around 1:1 as found in rabbit tissue [32].

Cell source and expansion
The post-mortem usage of biological material for biomedical research taken from the food chain requires no ethical permit according to the Swiss and European law. Porcine notochordal cells (NC) were isolated from the nucleus pulposus (NP) tissue of porcine tails obtained from 4-to 5-month-old pigs collected from the local abattoir. Tails with a high percentage of NC (~80%) were selected for the experiment. The percentage of NCs was assessed and counted using a haemocytometer and brightfield microscopy to confirmed the NC phenotype as large in cell size and presence of vacuoles. Bovine nucleus pulposus cells (NPC) were harvested from the NP tissue of~1-year-old bovine tails obtained from the local slaughter house. Porcine and bovine NP tissues were digested by 0.19% pronase (Roche, Basel, Switzerland) for 1 hour and subsequent collagenase type 2 at a concentration of 32 U/mL in 25 mL of HG-DMEM supplemented with 10% FCS (Worthington, London, UK) overnight (~14 hours) at 37°C and standard culture conditions (5% CO2 and 100% humidity) [14,23].
Cell separation using micro sieves and cytometry using FACS Cells isolated from porcine NP tissues were separated with three cell sieves, i.e., cells > 20 μm, 8-20 μm and cells < 8 μm (CellMicroSieves, BioDesign Inc., New York, USA) [33] with the goal to identify NPC and NC. To characterize and confirm the cell populations by size, the filtered and unlabeled cell fractions were resuspended in phosphate buffered saline (PBS) containing 0.5% bovine serum albumin (BSA) and were then analyzed on a fluorescenceactivated cell sorting (FACS) device (LSR II H271, Bdbiosciences, inc., Allschwil, Switzerland) with 429V for forward scattering (FSC-A) and 308V for side scattering (SSC-A). Gates were analyzed and visualized using FlowJo software 10.07R2 (TreeStar, inc., Ashland, OR).
Primary gates ( Figure 2) for small pNPC and bigger notochordal cells (pNC) were defined based on primary cells digested from discs lacking a notochordal cell phenotype, including bovine and a whole human lumbar IVD obtained from a spinal fusion (ethical permit has been obtained from the local ethical committee). The size of these cells was compared to the cell size of NC isolated from rabbit IVD and porcine IVD. Cells were grown in 2D monolayers for three passages or in 3D alginate beads at a density of 4M/mL alginate in Dulbecco's High Glucose Modified Eagle Medium (HG-DMEM, containing 4.5 g/mL glucose, Gibco, Lucerne, Switzerland) +10% FCS in order to test for cell size shifts during the cell expansion and to compare 2D monolayers with 3D alginate cultures.

3D cell encapsulation and co-culture
The cells were re-suspended in 1.2% alginate at a density of 4 M/mL and embedded in~30 μl beads by pressing the suspension at a constant rate through a syringe (22G needle) and dropping them into a 102 mM CaCl 2 salt solution [34]. The porcine NC:NPC ratio in the tissue was estimated to be~80% notochordal [23]. The cells were kept in co-cultures of NC: NPC ratios of 1:1 in serum-free HG-DMEM medium containing 100 μg/mL penicillin/ streptomycin, 50 μg/mL ascorbic acid, ITS+ and nonessential amino acids or in HG-DMEM supplemented with 10% fetal calf serum (all from Sigma-Aldrich, Buchs, Switzerland) ( Figure 1). All bead-bead co-cultures were conducted in duplicate in 12-well plates, using 0.4 μm pore size, high pore density, polyethylene terephthalate (PET) track-etched culture inserts (Becton, Dickinson and Company, Allschwil, Switzerland). The co-cultures were Figure 1 Experimental set-up to test NC regenerative effects on co-culture with NPC or AF that were stimulated initially in pre-culture with and without 10% fetal calf serum (FCS) and subsequent co-culture or exposure with conditioned medium (NCCM) from step 1. The design also involved a comparison of normoxia (~20% oxygen) and hypoxic conditions (2% oxygen).
tracked on day 0, day 7, and day 14. Day 0 was defined as the day of cell embedding into the 3D alginate environment. There were four co-culture pairings (thus, each N = 4 for the porcine and bovine animals) (see also Figure 1). The cultures were either kept in normoxia (20% atmospheric oxygen) or in hypoxia using a C-274-2-shelf chamber inside a standard incubator and 1x Pro-Ox controller, (Biospherix, inc., NY) and keeping O 2 at 2% by addition of N 2 .

Metabolic activity
Cell activity of the cells in alginate beads was measured using resazurin assay [35]. Two beads per condition were incubated in 500 μL DMEM with 10% FCS and 50 μM resazurin sodium salt (Sigma-Aldrich, Buchs Switzerland) for 3 hours in a 48-well plate [23]. The relative fluorescence unit (RFU) was then measured at an excitation wavelength of 547 nm and an emission wavelength of 582 nm using a Softmax pro multi-wave length fluorescence reader (Molecular Devices, distributed by Bucher Biotec, Switzerland). RFU values were expressed relative to the activity measured on day 0, i.e. the day of seeding the cells into alginate beads.

Quantification of GAG and DNA content
Alginate beads from the resazurin assay were digested with papain (Sigma-Aldrich, Buchs, Switzerland) overnight at 60°C. The samples digested with papain were used for glycosaminoglycan (GAG) and DNA measurement. The GAG content was measured by the modified dimethylmethylene blue (DMMB) assay (pH 1.5) [36,37]. The absorbance of the samples added to the DMMB buffer was read at 595 nm with a fluorescence reader (Molecular Devices). GAG concentrations were calculated from a standard curve obtained with chondroitin sulfate (Sigma-Aldrich, Buchs, Switzerland). The amount of DNA in the sample was measured with bisbenzimidol fluorescent dye (Hoechst 33258, Sigma-Aldrich). Fluorescence was again detected with a multi-wavelength fluorescence reader (Molecular Devices). A standard curve was generated with known concentrations of calf thymus DNA (Sigma-Aldrich, Buchs, Switzerland), and the amount of DNA of each sample was calculated from the standard curve.

Real-time RT-PCR
Relative gene expression of major anabolic genes of to the extracellular matrix turnover was monitored, i.e., aggrecan (ACAN), collagen type 1 and 2 (Col 1 and Col 2, respectively) and ribosomal 18S used as a reference gene [38][39][40]. Typical nucleus pulposus and notochordal cell markers including brachyury (T) and CD24 [41] were monitored. Due to DNA substitutions between the porcine and bovine codon sequences of one of the two primer regions, species-specific RT-PCR were designed (Table 1) for both NCs and NPCs. Relative gene expression was estimated by application of a threshold cycle (Ct) and calculation of ΔΔCt and the statistics of the 2-ΔΔCt according to Livak & Schmittgen [42].

Mass spectrometry analysis and protein identification of conditioned medium
Proteins of 1) serum-free control medium versus 2) NCCM and 3) medium collected from NPC and NC coculture were lysed in SDS-PAGE sample buffer and proteins separated on 10% SDS-PAGE gels. Gels were stained with Sypro Ruby® protein gel stain (Sigma), and the separated proteins were isolated from the gel according to Heller et al. [43]. Briefly, peptide sequencing was made on a LTQ Orbitrap XL mass spectrometer (ThermoFisher Scientific, Bremen, Germany) equipped with a Rheos Allegro nano flow system with AFM flow splitting (Flux Instruments, Reinach, Switzerland) and a nano electrospray ion source operated at a voltage of 1.7 kV. Peptide separation was performed on a Magic C18 column (5 μm, 100 Å, 0.075 × 70 mm) using a flow rate of~400 nL/min and a linear gradient of 5 to 40% acetonitrile in water/ 0.1% (v/v) formic acid over 60 min. Data acquisition was also done according to Heller et al. [43]. Using the Hardklör Database, mascot generic files were generated [44]. Applying a two round strategy, the searches were done with Phenyx [45] against the forward and reversed UniprotKB SwissProt protein database (Release 2013_12) of human entries. The parameters for the peptide search on the SwissProt protein database were also conducted according to Al Kaabi et al. [46]. The peptide match score summation (PMSS) value for each identified protein was used as a semi-quantitative abundance estimate. PMSS values were normalized by division with the median PMSS across all proteins in each sample and multiplication with the median across all samples. We use the SwissProt ID as the protein name in this manuscript.

Histology
The alginate beads were initially fixed in Tissue-Tek® O.C. T.™ compound (Sakura Finetek Europe, Leiden, The Netherlands) for 20 min and then snap-frozen in liquid nitrogen. Beads were then sectioned in 12 μm slices with a cryotome (Microm HM560, Thermoscientific, Reinach, Switzerland). For immune-staining, slices were incubated in solutions containing 0.1M CaCl 2 to prevent dissolution of beads. The cells were first permeabilized with 100% methanol for 2 min and then blocked with 10% FBS 0.1M CaCl 2 for 1 hour. The beads were then incubated with polyclonal rabbit brachyury (H-210) polyclonal primary antibody for 1h at a 1:200 dilution (sc-20109, Santa Cruz biotech, Santa Cruz, CA, USA). After washing, the primary antibody was visualized with the secondary antibody Alexa Fluor® 488 Goat anti-Rabbit IgG antibody (A1108, Molecular Probes, Invitrogen, Basel, Switzerland) at a 1:500 dilution. Sections were mounted in ProLong® Gold Antifade Reagent with DAPI (P-36931, Molecular Probes) to stain the nuclei and were visualized on an inverted fluorescent microscope (Nikon Eclipse E800, Nikon Instruments, Tokyo, Japan).

Statistical analyses
All data are given as relative to the "pure" cell population of the same culture day. "Pure" is defined as primary isolated cells cultured in 3D and no contact with other cells, i.e., the cells cultured in media lacking FCS and kept in normoxia. Statistical significance was tested using non-parametric Kruskal-Wallis exact test and multiple comparison tests using GraphPad Prism version 5.0f, GraphPad Software, San Diego California USA, www.graphpad.com.

Analysis of 2D and 3D cell populations
Cytometry was able to distinguish the cell size differences of the different cell types with the forward (F-SC) and side scatter (S-SC) (Figure 2). Initially putting cell isolations of different species unlabeled under the flow cytometer revealed the clear size differences of this primary isolated cell population after pronase/collagenase 2 digestion, with cell populations containing NC being much larger in F-SC and S-SC. This allowed the   definition of the major gate for NC based on cell populations from the mixed IVD cell type. The gate for NPC was based on cells isolated from IVDs lacking the NC fraction, such as bovine and human IVD ( Figure 2C, and D). Small NPC (< 8 μm cell size), a mixed cell population (8-20 μm), and a cell population with mainly large cells (>20 μm) were separated. Of the cells > 20 μm, almost all cells had a notochordal cell phenotype at the beginning of the culture. However, after 1-3 passages in 2D, the originally distinct cell populations turned into populations that were not distinguishable from each other ( Figure 3A). In contrast, much more NC could be recovered after 34 days of 3D alginate bead culture (around 50%, see Figure 3B). Thus, 3D alginate culture seemed to be superior for NC culture as a higher fraction of large NC were revealed after 34 days of culture, compared to the identical cell population cultured in 2D monolayer and with the requirement to passage them three times for the same number of culture days ( Figure 3).
Cell metabolism, proliferation, and GAG synthesis of 3D co-culture experiments   0.117) but was significant among the experimental groups in hypoxia (HY) after 7 days (K-W, p = 0.0163, Figure 4A). After 14 days, activity was not different among NC groups, neither under NO nor HY (for NO K-W, p = 0.322, for HY K-W, p = 0.051) ( Figure 4A and B). Pre-culture of NC prior co-culture revealed in general higher activity in the NC than in the controls after 7 days of co-culture. However, these effects flattened out after 14 days. NPC cell activity remained unchanged under NO after 7 days. There were no significant differences in cell activity among the NPC and the AFC experimental groups, whether they were cultured in NO or HY ( Figure 4E and F). The GAG/DNA ratio among the NC groups was significantly different after 7 days in both NO (K-W, p = 0.023, Figure 5A) and HY conditions (K-W, p = 0.016 for HY, Figure 5A), and the ratio remained significantly different after 14 days in HY (K-W, p = 0.044, Figure 5B). In NO conditions, the GAG/ DNA ratio of pure NPC was decreased on day 14 to 89.9 ± 38.3% (mean ± SEM) ( Figure 5D). When these NPC were co-cultured with NC 1:1, which were never exposed to FCS, the GAG/DNA ratio dropped after 7 days (i.e. 77.1 ± 18.68%, Figure 5C). However, in NO, if we added NCCM or co-cultured NPC with NC, which were exposed to 10% FCS during pre-culture, the ratio tended to be increased (190.6 ± 112.8%) or by addition of NCCM (140.1 ± 71.5%, Figure 5D) after 14 days. However, this was statistically non-significant among NPC groups (K-W, p = 0.964). In HY, however, the ratio tended to be elevated to 152.1 ± 50.2% for the CoC and to 195.7% ± 112.5% for the NCCM group ( Figure 5D). After 14 days of culture, these trends were maintained, but these were not significant.

Relative gene expression of NC
ACAN expression of NC was found up-regulated around ten times in the co-culture groups with NPC and previous exposure to FCS, compared to the pure cell population without previous contact with FCS, both in NO and HY ( Figure 6A). Further, we found that ACAN gene expression were more pronounced in the co-cultures supplied with 10% FCS and were more strongly expressed in the NO cultures ( Figure 6). Collagen type 2 expression was up-regulated much stronger in all groups cultured in NO but much less in HY. The gene expression of one of the major genes for the characterization of the notochordal cellsbrachyury (T)showed a weak up-regulation after 14 days of culture in the HY group, but not in the NO group ( Figure 6C).

Relative gene expression of NPC
RT-PCR data suggested that CD24 expression in bovine NPC (Figure 7), the NP-specific surface marker, was expressed differently in NO and HY conditions. While there was only a weak effect between the experimental groups of co-culture, however, the effect of the oxygen level was significant (K-W, p = 0.034). However, according to the ACAN/ col2 ratio criterion, the most "IVD-like" gene expression pattern among all investigated NPC groups was identified in HY and by co-culture with NC that were previously activated with FCS or by addition of NCCM (Figure 8).

Histology
The immune histology of NC in beads for brachyury (T) revealed a slightly stronger expression in the case of NO culture conditions compared to HY but the strongest effect was seen if NC were kept "boosted" in media containing 10% FCS for 7 days previous to co-culture ( Figure 9).

Mass spectrometry of NCCM
For the analysis of the medium protein profile (Table 2,  Additional file 1: Table S1), we screened for strong signal differences between the unconditioned and the conditioned medium (pooled after 7 days of serum-free culture). A full table of the Swiss Prot search of identified proteins can be obtained as an additional supplemental, Additional file 1: Table S1. We found that the CTGF increased in the conditioned medium, which has been identified in canine notochordal cells by Erwin et al. [24]. Further, we found a high concentration of the serum amyloid A-2 protein (SAA2_MUSVI), an outbreak protein identified in immune burst reactions (Table 2 and Additional file 1: Table S1).

Phenotype of NC
The NC phenotype is specific in the sense that cells are relatively large in size [47] and highly dependent on nutrition gene expression bovine NPC because of a high cellular activity [23,28]. Here, we present data that clearly demonstrate that in monolayer cell culture, NC seem to be outcompeted by the small chondrocyte-like NPC (Figure 3). Of course, we cannot conclude from our unlabeled FACS cytometric data whether the NC differentiated into NPC or whether NPC simply grew faster than NC. However, a recent time-lapse microscopy study by Kim et al. [48] showed that NC populations harvested from the spine of mature New Zealand white rabbits proliferate much slower than the smaller chondrocyte-like cells (i.e. defined as NPC in our case). These authors, furthermore, demonstrated, using time-lapse cell-tracking microscopy, that by looking closer into the NC fraction, there are different cell types present, i.e., vacuolated, giant and polygonal cells that differ in cell proliferation and speed of cell motility. These authors suggested that NC differentiated in vitro into these three morphologically different cell types. The negative effects of monolayer culture for the maintenance for NC  has also been demonstrated by Rastogi et al. [26], who cultured rat NC in monolayer in NO and HY. They found that 3D alginate culture maintained the NC phenotype better compared to 2D monolayer. The rapid loss of NC over time can be partially prevented by 3D alginate culture for up to 34 days, as was revealed in this study (Figure 3). Considering our data and the published work, it is still unclear which scenario might be true. We further confirm a strong nutrient dependency (by addition or lack of FCS), as has been found previously [28]. We can clearly confirm a strong dependency of NC to culture medium composition and also oxygen content, as revealed by cell activity and GAG/DNA ratio data (Figures 4 and 5). We found a trend towards increased GAG/DNA ratio if NC were kept in medium containing 10% FCS ( Figure 5). It has been demonstrated that NC and NPC differ in cell size, nutrition [28], surface markers [49], and mechanosensitivity [29]. It was found that NC glucose consumption rate is higher than in NPC under identical culture conditions [28]. Furthermore, the clear size differences were confirmed by non-invasive femtosecond laser microscopy [47]. Furthermore, the NC differ by the presence of large vacuoles, which can be separated by the size-scatter of FACS analysis [3,6], as has also been shown by our study. These large vacuoles found in NC of the intervertebral disc have been attributed to a possible functional role in Table 2 Summary of mass spectrometric analysis of the most interesting proteins related to the IVD extracellular matrix homeostasis identified in serum free medium (SFM) on day 0, conditioned medium (NCCM) of pure NC collected after 7 days and co-culture (CoC) medium of 1:  osmoregulation [50]. It has been noted that these vacuoles are not present anymore in the NC after long-time in-vitro culture of 28 days using porcine NC [27,51]. No such complete loss of vacuoles but a reduction in numbers were reported by other in vitro studies using rat or rabbit NC [26,48]. Transcriptomics that compare the NC and the NPC cell population revealed that there are only about 20 genes really distinct between these two cell types [10,52]. A number of cell surface markers that separate these two cell populations have been identified [49]. Weiler et al. [53] found that cells in the human fetal and juvenile NP with the typical morphology of the notochord (physaliferous) express the markers cytokeratin CK-8, -18, -19, and Galectin-3 [54]. Another NC-typical marker is brachyury (T), which was investigated in this study. We found that the notochordal homeobox gene marker T did not change when altering the culture conditions but decreased about 10-fold under normoxia at the RNA level ( Figure 6). Immunohistology staining confirmed this finding and showed a slight increase of T expression in the case of hypoxia ( Figure 9). Potier et al. [27] found a general loss of the NC phenotype and loss of the markers cytokeratin 8 and 19 (KRT8,19) in their co-culture system of NC, NPC, and mesenchymal stem cells over 28 days of culture. Gilson et al. [49] found that pig NPC, which are phenotypically similar to human infant NPC, were all KRT8 positive. This has been recently confirmed by another histological study on human tissue; the authors furthermore demonstrated that KRT8 is down-regulated with onset of IVD degeneration [55]. However, as we did not look into KRT8 expression in this study, we cannot comment on the validity of the KRT8 marker. Interestingly, a strong effect between hypoxic and normoxic condition was found in the surface marker CD24 in the NPC (K-W test, p = 0.034) (Figure 7), such that CD24 was higher expressed under HY. This finding is in agreement with the culture study presented by Rastogi et al. [26] where they kept NC in alginate beads and in hypoxia and normoxia. Thus, CD24 might indeed represent a sensitive marker for the NC phenotype. CD24, a glycosylphosphatidylinositol (GPI) anchor protein, has been proposed as a NP-specific marker [26,41]. Finally, a Tie2+ cell population has been recently reported about, which reflects a stem cell population in the IVD, which decreases rapidly with age [56]. These cells, as well as induced progenitor (iPS) cells, seem to express CD24 and can be enriched via this surface marker [16,56]. iPS cells, which hold potential for the regeneration of the IVD, could at least theoretically be produced from both NC and NPC. However, it is still unclear which cell type would be more potent and suitable.
Optimizing NC culture conditions NC are perfectly adapted to a low oxygen environment. They can produce a better-aligned ECM under hypoxic conditions [20]. Here, we cultured the cells under normoxic and hypoxic conditions. We found that the GAG/ DNA ratio of the NC could be significantly increased indeed under HY rather than NO after 14 days ( Figure 5B). However, in terms of the better maintenance of a notochord phenotype, the expression of T was higher under HY than in NO from both gene expression and immunestaining data. It could also be that keeping the NC in clusters rather than completely isolated by digestion and increasing the osmolarity from 230 mOsm/L to 400 mOsm/L could preserve the porcine NC phenotype better in culture [51]. Interestingly, we found that by applying indirect co-culture or applying the CM to the bovine NPC, we could obtain a ACAN/col 2 ratio, which was most favorable for the IVD phenotype, in the order of a 1:1 ratio as found in rabbit cells [32] (Figure 8). Similar findings were obtained by previous studies culturing NC or NPC under hypoxia [16,20,31,57]. We cultured the NC in high glucose DMEM (i.e. with 25 mM glucose) either in serum-free medium or medium supplemented with 10% FCS. Park and Park [58] found that the cell proliferation of NC was significantly decreased at glucose concentrations of 100 mM, 200 mM, and 400 mM and apoptosis was increased via the intrinsic pathway with dose-and time-dependent effects. Guehring et al. [28] tested the nutrient sensitivity of NC depleting glucose from 5 mM to 0 mM and found a strong nutrient sensitivity of these cells. Future experiments could be to test culture conditions on NC viability in the 5-20 mM range and adjusting osmolarity.

Composition of NCCM
The analysis of the NCCM by mass spectrometry revealed the presence of expected anabolic proteins, such as the collagens 1, 5, and 6, which these cells produce in 2D and surely in 3D culture (Table 2). Notably, collagen type 2 was not hit by the SwissProt blast. There were also two MMPs detected (MMP1 and MMP2) detected, which is not unexpected given the nature of the NC and NPC. However, unexpectedly, neither MMP3 nor MMP13 were hit by the SwissProt search in the NCCM. These missed proteins might be due to the limited approach to analyzing the NCCM and CoC-medium presented here. Future experimental work could specifically isolate the vacuoles of the NC through centrifugation methods and then perform mass spectrometry analysis on these fractions, which might reveal more focused spectrum searches. Another interesting candidate found by the SwissProt search was High-Temperature Requirement A Serine Peptidase (HTRA1) that was present in the case of the 7 days CoC of NC with NPC (Table 2, Additional file 1: Table S1). This enzyme has been described as a potential key enzyme in intervertebral disc


s2fieldsofstudy: Biology Medicine

text:
Cytokine secretion from brain macrophages infected with human immunodeficiency virus in vitro and treated with raltegravir

Background Integrase inhibitors are a promising class of antiretroviral drugs to treat chronic human immunodeficiency virus (HIV) infection. During HIV infection, macrophages can extravasate from the blood to the brain, while producing chemotaxic proteins and cytokines, which have detrimental effects on central nervous system cells. The main goal of this study was to understand the effects of raltegravir (RAL) on human brain macrophage production of immune-mediators when infected with HIV, but did not compare with other antiretroviral agents. Methods Pro-inflammatory cytokines, IFN-γ, IL-10, IL-12-p70, IL-1, IL-8, TNF-α, and IL-6 were measured simultaneously in tissue culture supernatants from primary brain derived macrophages, microglia. We tested the effects of RAL on markers of astrocytosis and neurite integrity in primary human neuroglial cultures. Results RAL administered at 20 nM effectively suppressed HIV infection in microglia over 9 days. Only IL-8, IL-10, and TNF-α were above the detection limit in the majority of samples and RAL significantly suppressed the rate of cytokine production in HIV-infected microglia. During RAL-alone, the rate of IL-8 secretion was higher. Conclusions RAL did not affect neurite area but inhibited astrocyte growth in the neuroglial cultures. Exploring the effects of RAL on pro-inflammatory molecule production in brain macrophages may contribute to designing ARV neuroprotective strategies in chronic HIV infection.

Background
Chronic human immunodeficiency virus (HIV) infection has detrimental effects on central nervous system (CNS) neuronal health. The principal targets of productive HIV infection in the brain are microglia and monocyte derived macrophages [1][2][3][4]. In the periphery, HIV infection in CD4 T cells is cytopathic, resulting in fast CD4 T cell depletion [5], whereas in resting memory T cells, tissue monocytes, macrophages, and brain microglia with integrated HIV survive longer and serve as an HIV reservoir that persists indefinitely [6]. In adult humans, parenchymal microglia are infrequently exchanged with peripheral blood monocytes [7]. Perivascular cells in the CNS with macrophage markers have a higher turnover and are more frequently replaced by blood monocytes. Therefore, perivascular brain macrophages may serve as a source of HIV in the brain, and immunohistological and DNA hybridization studies have confirmed the presence of HIV proteins and DNA in these cells [8], which has been shown to be associated with depleted pre-and post-synaptic markers, synaptophysin and microtubule associated protein-2 (MAP2) [9].
HIV effects on the CNS have behavioral consequences. Neurocognitive impairment in HIV infected individuals is associated with elevated viral load and microglial activation [10,11]. There are several prevailing hypotheses for the mechanisms of neural damage in HIV: direct neurotoxic effect from viral proteins and neural damage via intracellular signaling and secondary neural damage from cytokines and chemokines from infected microglia and brain macrophages [12]. Drugs used to treat HIV have variable neurotoxic effects, as examined by MAP2 staining in neuronal cultures, ranging from 17% to 52% reduction in MAP2 area. Of six combination regimens which included protease inhibitor, non-nucleoside reverse transcriptase inhibitor, and reverse transcriptase inhibitor, four caused significant reduction in neuronal MAP2 area [13]. As new antiretroviral drugs become available, it is important to continue assessing neurotoxicity.
Integrase inhibitors are a relatively new class of antiretroviral compounds, one of which is raltegravir (RAL), which has potent and durable antiviral activity similar to that of efavirenz, but which achieved HIV suppression to below detectable limits at a faster rate in a 24 week study [14]. HIV integration into the host genome is a multistep process involving: a) 3′ endonucleolytic cleavage of the 3′ end of DNA in the HIV pre-integration complex to a conserved CA dinucleotide, b) Strand-transfer involving concerted cleavage of host DNA and ligation of viral 3′ DNA to 5′ staggered site on host DNA, and c) DNA repair of gaps and DNA synthesis. RAL functions by inhibiting the strand-transfer step [15], thus preventing the induction of DNA repair machinery which is associated with innate immune activation and cell death [16]. We were therefore interested in determining the CNS efficacy of RAL by assessing: 1) neurotoxicity using in vitro neuronal cultures following methods comparable to a previous study [13], 2) inhibition of HIV infection in cultured microglia, and 3) cytokine production in RALtreated, HIV infected microglia.
In primary human neuroglial cultures, we found that RAL is not neurotoxic and we present evidence for reduced astrogliosis. In primary human HLA-DR-positive brain macrophages and microglia, we measured secretion of seven cytokines across 9 d of in vitro HIV infection, and found reduced rate of TNF-α, IL-10, and IL-8 production in those treated with 20 nM RAL.

Cell culture
This study was approved by University of California San Diego Human Research Protections Program. Microglia were isolated through differential adhesion procedure from fetal human brain tissue from elective terminated pregnancy between 12 and 16 weeks of gestation [17], acquired from Advanced Bioscience Resources, donors gave written informed consent for research-use of the cells and tissue. Single-cell suspension of central nervous system cells were plated at 10 8 cells/mL in a 125 cm 2 flask and after 7 d, microglia removed by agitation and withdrawal of nonadherent cells which were plated in selection media on coverslips coated with poly-L-lysine (Life Technologies P4707, Carlsbad, CA, USA). Microglia media components were purchased from Life Technologies and consisted of DMEM (11965-092), supplemented with glutamax (35050-061), and gentamicin sulfate (15710-064). After 4 d in culture, cells were differentiated with granulocyte/ macrophage colony stimulating factor (GM-CSF, Fisher Scientific, 5056909) for 1 d. Thereafter, no media contained GM-CSF. Microglia were inoculated with a seed stock of HIV Ba-L (5,000 pg/mL HIV p24) in fresh media for 4 hr, then media changed in the presence and absence of 20 nM RAL and maintained without media changes for 9 d. The following reagents were obtained through the NIH AIDS Reagent Program, Division of AIDS, NIAID, NIH: HIV-1 Ba-L from Dr. Suzanne Gartner, Dr. Mikulas Popovic and Dr. Robert Gallo [18] and RAL (Cat # 11680) from Merck & Company, Inc. All experimental conditions were performed with three independent replicates.
Aliquots of 60 μL supernatant were removed every 48 hr starting 1 d after infection and stored with 2.5 μL 25X Complete protease inhibitor (Roche 04693116001, Indianapolis, IN, USA) and stored at −80°C to measure cytokine production. The 60 μL was replaced with fresh media (no GM-CSF supplemented) to maintain a constant 500 μL per well. To verify HIV infection, a 200 μL aliquot of supernatant was removed and replaced with fresh media for HIV p24 ELISA at 1 d of infection and at the endpoint (9 d) (Advanced Bioscience Laboratories, Inc. 5421, Rockville, MD, USA). After 9 d, cells were fixed in 4% paraformaldehyde (Electron Microscopy Sciences 15710-S, Hatfield, PA, USA) for microscopy.
To assess for cytotoxicity, 25 μL supernatant at endpoint (9 d) was used to measure lactate dehydrogenase (LDH) activity following manufacturer's instructions of CytoTox 96 Non-radioactive Cytotoxicity Assay (Promega, Madison, WI, USA). Percent-cytotoxicity was calculated for each condition by comparing to LDH activity of supernatant from cells lysed in 0.1% Triton-X 100 for 1 hr.
Neuronal cultures were generated according to a protocol first described by White et al. [19] and modified for our current usage exactly as described by Nguyen et al. [20] from donated fetal human brain tissue from elective terminated pregnancy between 12 and 16 weeks of gestation. Single-cell suspension of central nervous tissue were plated at 10 5 cells/mL on glass coverslips coated in poly-L-orinithine and laminin in 24 well-plates. Cultures were mixed neuron -glia culture composed of approximately 60% neurons and 40% astrocytes. After 28 d in culture, neurons were exposed to 20 and 100 nM RAL overnight following by fixation and microscopy analysis.

Microscopy
Cells were fixed in 4% paraformaldehyde for 10 min at 37°C and washed 3 times in Tris buffered saline (PBS). Cells were permeablized and non-specific antigen blocked using 0.2% Triton X-100 and 2% fetal bovine serum for 2 hr, then incubated overnight with primary antibody at 4°C. After washing 3 times in TBS-T (0.2% Tween-20), secondary antibodies were incubated at 1:750 dilution in blocking buffer, then washed 3 times in TBS-T, a final wash in distilled water, then finally mounted on glass slides with Vectashield with DAPI mounting media (Vector Labs H-1500, Burlingham, CA, USA). Images were captured by laser scanning confocal microscopy at 40X magnification for quantification, images were captured for 10 random fields per coverslip. The imaging conditions were maintained exactly the same across the different experimental conditions (separately for measuring β-III tubulin, GFAP, or NF-κ-B). The following primary antibodies were used, diluted in blocking buffer: mouse anti β-III tubulin (1:400)  For image quantification of neuronal cultures, methods were adapted from White et al. [19], green (β-III tubulin) and red (GFAP) channels were separately measured by setting a signal threshold and determining the area covered, representing neurite density for β-III-tubulin, and astrogliosis for GFAP. To quantify nuclear translocation of NF-kB, methods were adapted from Tatro et al. [21], the cells were delineated by creating a mask for the HLA-DR signal representing the cell bodies and the nuclei were delineated by creating a mask for the DAPI signal. The NF-kB signal was calculated for the cell body and the nuclei by total signal intensity, and then the percentnuclear and percent-cytoplasm were calculated.

Statistical analysis
For GFAP and β-III tubulin comparisons, Student's t-test was used to compare RAL vs Control of area covered. For NF-kB measurements, percent nuclear was arcsinetransformed (to account for upper and lower limits, 0-100%) and each condition compared by Student's t test. For cytokine production, linear regression using Least Squares of concentration vs. time for each condition (Control, HIV, RAL, HIV + RAL) was calculated (fitted to Equation 1), including an interaction term. Equation 1: [Cytokine] = β 0 + β 1 Time + β 2 Condition + β 3 Condition*-Time, we tested for effects of time (rate of secretion), main effect of Condition, and a Condition × Time interaction. β 3 corresponds to the effect that a given condition has on the cytokine secretion compared to the null hypothesis, P-values reported to test for a Condition × Time interaction is the probability that rate is the same for all conditions together. The rates of cytokine production (pg/mL⋅d) for each treatment condition were compared to Control by Dunnett's test.

Results and discussion
In this study, we measured cytokine secretion from human microglia under four conditions, 1) No-treatment control, 2) Infected with HIV, 3) Infected with HIV but treated with 20 nM RAL, and 4) 20 nM RAL alone. The concentration of 20 nM RAL was chosen on the basis of evaluating several pharmacokinetics studies. The mean trough plasma concentration from RAL once-daily parmacokinetics studies was 40 nM [22], while an independent study calculated a CSF:Blood Plasma ratio of 0.058 with a median (over 24 hr) plasma concentration 540.7 nM (31.36 nM in CSF) [23]. One additional study evaluated intracellular RAL concentration and calculated a 24 hr area under the curve (AUC) 1,884 nMxh [24], which is averages to roughly 78.5 nM. Therefore 20 nM RAL seemed to be a reasonably relevant concentration to assess in vitro effects on microglia.
We found that RAL administered at 20 nM was effective at suppressing HIV infection in microglia (Figure 1) for at least 9 d. Only IL-8, IL-10, and TNF-α were above the detection limit. We calculated the rate of cytokine production for all three cytokines across the different treatment groups ( Table 1). The mean IL-8, IL-10, and TNF-α concentrations and linear regression, separated by conditions are shown in Figure 2. For IL-10, IL-8 and TNF-α, there was a significant effect of RAL on the cytokine production in the context of HIV infection. Alone, the RAL-treated microglia had the highest concentration of IL-8, IL-10, and TNF-α. However, in the context of HIV infection, RAL-treated microglia had lowest production of TNF-α and IL-8. This makes sense because TNF-α autocrine signaling leads to IL-8 production via NF-kB. However, it is important to note that the highest production of TNF-α was in the presence of RAL alone (Table 1). IL-8 and IL-10 production were lowest in RAL treated microglia in the context of HIV, but not significantly different among the other groups (Table 1).
It is remarkable that HIV and RAL alone induced cytokine expression, but in combination, was below control levels. This implies that RAL may be pro-inflammatory alone, and in the presence of HIV replication complex it is anti-inflammatory. One possibility is off-target effects on DNA-binding proteins, mimicking NF-kB -like activation, however in the presence of HIV-replication complex, it is bound and not interacting with off-target proteins.
We assessed cytotoxicity through measuring LDHrelease in HIV-infected cells exposed to RAL and found background levels cytotoxicity due to HIV-infection, which was neither enhanced or diminished by RAL. The mean ± standard deviation cytotoxicity in HIV-alone was 43 ± 7.9%; and for HIV + RAL was 45 ± 3.2%. This may have been due to the protocol, which was acute infection via exposure to high levels of HIV for 4 hr, followed by media change including the drug. RAL may have suppressed further integration and infection of more cells with HIV, but it did not prevent lysis/death of those already infected.
Activation of the IL-8 gene is enhanced by signaling from NF-kB, and based on reduced IL-8 secretion in the HIV + RAL treated cultures, we hypothesized lower NF-kB activation and quantitated the nuclear translocation of NF-kB after 9 d infection, we calculated the percentnuclear NF-kB compared to total NF-kB. The proportion of NF-kB localized to the nucleus was significantly lower in the HIV + RAL than the HIV + alone. Nuclear localization of phosopho-NF-kB was quantitated as percent-nuclear, then arcsine transformed for comparison by Dunnett's test vs. Control. RAL alone (21%) and HIV + RAL (22%) were not significantly different from Control (20%), while HIV + alone was significantly higher from Control (28%, P = 0.02).
There are several cellular markers of neurons and astrocytes in cell culture, one of which is class-III βtubulin, a structural microtubule protein specifically expressed in the cell body, axon, and dendrites of neuronal cells. To determine the neurotoxicity, we measured the area of coverage of β-III tubulin, similar to Robertson et al. [13], because retraction or loss of neurites would be detected by this method. In order to measure the relative amount of astrocytes in the culture, we similarly measured the area covered by the astrocyte-specific protein, GFAP. Diffuse astrocytosis was observed during HIV associated dementia and HIV-encephalitis [25], and in the CNS of transgenic mice expressing HIV proteins [26] and is a marker of astrogliosis or astrocytosis. In the absence of RAL, neuronal cultures had 110 ± 41 μm 2 β-III tubulin and 244 ± 49 μm 2 GFAP. While with 20 and 100 nM RAL, there was no significant effect on β-III tubulin, with 142 ± 48 and 89.4 ± 34 μm 2 , respectively, with P = 0.17 and P = 0.43 as assessed by Dunnett's test vs. Control. However, with 20 and 100 nM RAL, there was a significant effect on GFAP, with 107 ± 26 and 115 ± 163 μm 2 , respectively, with P = 0.0094 and P = 0.01 as assessed by Dunnett's test vs. Control. Thus, RAL had no significant effect on β-III tubulin area at 20 or 100 nM and an inhibitory effect on GFAP area. The possibility that RAL is toxic to astrocytes should be noted, considering the important role that they play in maintaining the blood brain barrier and neuronal maintenance. Figure 3 shows the quantitation of β-III tubulin and GFAP area as well as representative photomicrographs.
The CNS has been proposed as a compartment for a latent HIV reservoir which has long-term neurological effects with downstream neurocognitive consequences, attributed to cytokine production by infected perivascular CNS macrophages and parenchymal microglia [10]. Therefore, understanding whether antiretroviral compounds themselves are neurotoxic becomes important when infected individuals are on therapy for decades. The purpose of the present study was to assess whether RAL, an inhibitor of strand-transfer, has neurotoxic properties and whether RAL slows the rate of cytokine secretion by HIV infected microglia. We found that RAL is not neurotoxic at 20 or 100 nM and that it significantly inhibits cytokine secretion in HIV infected microglia in vitro.   Robertson et al. [13] found that exposure of primary human neurons caused statistically significant reduction of MAP2, a dendritic marker of mature neurons, when exposed to some antiretroviral drugs. Individually, abacavir, 2′,3′-dideoxyinosine, and nevirapine are predicted to have relatively high risk of neurotoxicity, with at least a 10% drop in MAP2 intensity at observed typical cerebrospinal fluid (CSF) concentrations of the drugs [13]. Median RAL concentration in the CSF of treated HIV patients was observed to be 14.5 ng/mL (30.05 nM) [23], which is within range of the concentrations we tested for neurotoxicity. This suggests that RAL has relatively low neurotoxic risk.
We measured cytokine production by withdrawal of supernatant from infected microglia cultures and quantification using a multiplex assay and found production of IL-8, IL-10, and TNF-α, which increased linearly. RAL on its own caused significant increase in the rate of production of these three cytokines, mainly IL-10, an anti-inflammatory cytokine; but resulted in a significant decrease when administered with HIV. The most abundant was IL-8, a molecule with potent chemotaxic properties for neutrophils [27] and monocytes [28]. IL-8 gene transcription is induced by NF-kB activation and nuclear translocation, and is dependent on TNF-α. One preliminary positron emission tomography (PET) study found slightly increased retention of the PET ligand 11 C-PK11195, which binds to activated microglia, in neurologically asymptomatic and ARV-treated HIV-infected individuals [11]. One human study assessed IL-8 concentration in CSF of patients with HIV associated dementia compared to HIVinfected patients with no neurocognitive impairment and found higher IL-8 in CSF of those with dementia [29]. In our experiments, phospho-NF-kB nuclear translocation was significantly higher only in the HIV-infected cultures, not in Control, RAL-alone, or HIV + RAL.
We also observed production of IL-10, an anti-inflammatory cytokine which inhibits T cell proliferation and is putatively produced early in HIV infection via NF-kB signaling [30], which would be an antiviral action, allowing the incipient reservoir to avoid detection.
In comparison with nucleoside reverse transcriptase inhibitors, tenofovir and zidovudine, RAL has relatively modest effects on monocyte cytokine production. Zidovudine dose-dependently increased secretion of IL-8 and tenofovir decreased it. Likewise, the anti-inflammatory IL-10 was reduced in the presence of tenofovir while zidovudine did not affect it [31].
One possible explanation for reduced TNF-α, IL-8, and IL-10 production in our cultures is simply due to fewer infected cells. Another possible explanation is that inhibition of the strand transfer step of HIV integration prevents the DNA damage and repair process from being initiated, which may lead to downstream microglial activation. One weakness to the present study is the lack of comparison with another antiretroviral compound or lack of testing for combinations. Additionally, with measuring the cytokines at one-day intervals, it is difficult to determine the order of events except by interpolating from what is already known. An important future experiment would be to assess the effect of conditioned media from brain macrophage cultures on neuronal and glia cultures.

Conclusions
Results from this study suggest a low probability for neurotoxicity of RAL and likely neuroprotective effect in HIV-infection by suppressing the production of chemotaxic inflammatory cytokine, IL-8.
(See figure on previous page.) Figure 2 IL-8, IL-10, TNF-α, IL-6, IFN-γ, IL-1β, and IL12p70 production in supernatant of microglia during 9 d in culture. Microglia were infected with HIV or not, and exposed to 20 nM RAL immediately afterwards (RAL, HIV + RAL), and a non-treated culture from the same source grown alongside (Control). Supernatants were withdrawn and cytokines measured by Mesoscale Discovery 7-plex pro-inflammatory cytokine kit. Plotted are concentrations (pg/mL) vs. Time after infection, separated by treatment group, error bars indicate standard deviation of biological triplicates, dotted horizontal line indicates detection limits. Based on Standard Least Squares linear regression, RAL-treatment in HIV-infection significantly reduced the secretion rate of IL-8, IL-10, and TNF-α. In RAL-alone, there was higher TNF-α and IL-6 at day 8. Figure 3 Primary human neuron-glia cultures exposed to RAL. (A) Quantification of area of β-III-tubulin (average area, μm 2 ), and GFAP in neuron -glia cultures exposed to 0, 20, and 100 nM RAL for 15 hr. Representative images of cultures treated with (B) Control, (C) 20 nM, and (D) 100 nM RAL. Cultures were maintained for four weeks in neurobasal media on glass coverslips then exposed to RAL overnight, cells were fixed and stained for β-III tubulin (green) and GFAP (red) to illustrate neurons and astrocytes, respectively. Ten images were acquired by deconvolution fluorescence microscopy under 40X objective magnification and quantified using Image Pro Plus.